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Musca Domestica L. (Diptera

Permanent Link: http://ufdc.ufl.edu/UFE0041965/00001

Material Information

Title: Musca Domestica L. (Diptera Muscidae) Dispersal from and Escherichia coli O157:H7 Prevalence on Dairies in North-Central Florida
Physical Description: 1 online resource (215 p.)
Language: english
Creator: Burrus, Roxanne
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2010

Subjects

Subjects / Keywords: bacteria, dairy, disease, emerging, enteric, escherichia, farm, fly, house, housefly, interface, muscid, pathogen, prevalence, rural, transmission, urban
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: House fly, Musca domestica L., dispersal up to 3 km from a dairy release site into a nearby town was documented in this study. Dispersal occurred by both direct flight across multiple, mixed habitat types including open fields with interspersed tree copses, and by corridors and edges provided by local roads. Marked flies were collected at an adjacent dairy, and at most traps that were set along two well-travelled roads connecting the dairies and the town. Additionally, one marked fly was captured on a trap that was placed outside of a restaurant in town 3 km from the release site. In total, 0.67% (250) of marked house flies were recaptured from a total release of 37,2000 marked flies over 11 wk. Escherichia coli O157:H7 was isolated from two dairies in north-central Florida using immunomagnetic separation followed by direct culture plating onto CHROMAgar and sorbitol MacConkey agar supplemented with potassium tellurite and cefixime (CT-SMAC) selective media. Presumptive identification of E. coli O157:H7 was confirmed by polymerase chain reaction (PCR) using the fliCH7 and rfbEO157 gene amplification. Dairy samples that were tested included pools of house flies, spilled grain, and fresh dairy cattle manure. Forty-two percent (24/57) of samples were positive by direct-culture using both CHROMAgar and CT-SMAC agar, with 11 positive samples from CHROMAgar and 13 positive samples from CT-SMAC agar. The 24 positive samples were submitted to PCR; of those 24 samples, 14 (58%) were confirmed by PCR. Two of the PCR-confirmed samples were false-negatives on CHROMAgar media, but presumptive positive on CT-SMAC media, indicating the importance of analyzing samples by more than one method, and demonstrating the sensitivity of PCR. Direct culture prevalence rates from CHROMAgar were 14.0% (8/57) for house flies, 5.3% (3/57) for grain, and 0% (0/57) for manure. The rate of CHROMAgar isolation of E. coli O157:H7 from house flies was 2.6 times greater than from grain. The PCR confirmation rates were 67% (8/12) for house flies, 56% (5/9) for grain and 33% (1/3) for manure. These data suggest that detection of E. coli O157:H7 on dairies might be more accurately determined by testing house flies instead of grain or manure samples, regardless of which isolation method is utilized. Flies are easy to collect and process, and because they disperse into urban areas, they provide valuable information regarding a mobile element for pathogen transmission that is lacking in grain and manure samples. House flies should be an important consideration in the design of a pathogen monitoring program on dairies.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Roxanne Burrus.
Thesis: Thesis (Ph.D.)--University of Florida, 2010.
Local: Adviser: Hogsette, Jerome A.
Local: Co-adviser: Kaufman, Phillip Edward.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2011-08-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2010
System ID: UFE0041965:00001

Permanent Link: http://ufdc.ufl.edu/UFE0041965/00001

Material Information

Title: Musca Domestica L. (Diptera Muscidae) Dispersal from and Escherichia coli O157:H7 Prevalence on Dairies in North-Central Florida
Physical Description: 1 online resource (215 p.)
Language: english
Creator: Burrus, Roxanne
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2010

Subjects

Subjects / Keywords: bacteria, dairy, disease, emerging, enteric, escherichia, farm, fly, house, housefly, interface, muscid, pathogen, prevalence, rural, transmission, urban
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, Ph.D.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: House fly, Musca domestica L., dispersal up to 3 km from a dairy release site into a nearby town was documented in this study. Dispersal occurred by both direct flight across multiple, mixed habitat types including open fields with interspersed tree copses, and by corridors and edges provided by local roads. Marked flies were collected at an adjacent dairy, and at most traps that were set along two well-travelled roads connecting the dairies and the town. Additionally, one marked fly was captured on a trap that was placed outside of a restaurant in town 3 km from the release site. In total, 0.67% (250) of marked house flies were recaptured from a total release of 37,2000 marked flies over 11 wk. Escherichia coli O157:H7 was isolated from two dairies in north-central Florida using immunomagnetic separation followed by direct culture plating onto CHROMAgar and sorbitol MacConkey agar supplemented with potassium tellurite and cefixime (CT-SMAC) selective media. Presumptive identification of E. coli O157:H7 was confirmed by polymerase chain reaction (PCR) using the fliCH7 and rfbEO157 gene amplification. Dairy samples that were tested included pools of house flies, spilled grain, and fresh dairy cattle manure. Forty-two percent (24/57) of samples were positive by direct-culture using both CHROMAgar and CT-SMAC agar, with 11 positive samples from CHROMAgar and 13 positive samples from CT-SMAC agar. The 24 positive samples were submitted to PCR; of those 24 samples, 14 (58%) were confirmed by PCR. Two of the PCR-confirmed samples were false-negatives on CHROMAgar media, but presumptive positive on CT-SMAC media, indicating the importance of analyzing samples by more than one method, and demonstrating the sensitivity of PCR. Direct culture prevalence rates from CHROMAgar were 14.0% (8/57) for house flies, 5.3% (3/57) for grain, and 0% (0/57) for manure. The rate of CHROMAgar isolation of E. coli O157:H7 from house flies was 2.6 times greater than from grain. The PCR confirmation rates were 67% (8/12) for house flies, 56% (5/9) for grain and 33% (1/3) for manure. These data suggest that detection of E. coli O157:H7 on dairies might be more accurately determined by testing house flies instead of grain or manure samples, regardless of which isolation method is utilized. Flies are easy to collect and process, and because they disperse into urban areas, they provide valuable information regarding a mobile element for pathogen transmission that is lacking in grain and manure samples. House flies should be an important consideration in the design of a pathogen monitoring program on dairies.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Roxanne Burrus.
Thesis: Thesis (Ph.D.)--University of Florida, 2010.
Local: Adviser: Hogsette, Jerome A.
Local: Co-adviser: Kaufman, Phillip Edward.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2011-08-31

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2010
System ID: UFE0041965:00001


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MUSCA DOMESTIC L. (DIPTERA: MUSCIDAE) DISPERSAL FROM AND ESCHERICHIA
COLI 0157:H7 PREVALENCE ON DAIRIES IN NORTH-CENTRAL FLORIDA




















By

ROXANNE GRACE BURRUS


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2010



































2010 Roxanne G. Burrus


































To family and friends









ACKNOWLEDGMENTS

Thanks to the following individuals who made completion of this dissertation possible.

Thanks to my committee members, Jerome A. Hogsette, Phillip E. Kaufman, James E. Maruniak,

Volker Mai, and Amarat Simonne. Their constructive suggestions provided insight and were

tremendously beneficial at many levels. Thanks to Heather Furlong, Lois Wood, Chris Geden,

and Melissa Doyle for assistance with the fly dispersal study and house fly rearing. Thanks to

Richard Robbins, Marianne Radziewicz and David Hill of the Armed Forces Pest Management

Board (AFPMB), for assistance with the literature. Thanks to Wei-Yea Hsu, Luis Mendoza,

James Becnel, and Julia Pridgeon, for assistance with microbiological processing of samples.

Thanks to Dongyan Wang for statistical assistance. Thanks to James E. Maruniak and Alejandra

Garcia-Maruniak, Michael Scharf, Volker Mai, and Maria for assistance with polymerase chain

reaction assays. Thanks to Debbie Hall, Katherine Smitherman and Mrs. Quispe for "over-the-

top" administrative support. Thanks to Navy Entomology personnel for mentoring and guidance,

including Stanton Cope, Gregory Beavers, George Schoeler, David Hoel; and from the Air

Force, Doug Burkett. On a more personal note, the following family members and friends

provided timely support, advice, encouragement, and inspiration: Dawn Burrus, Karen Burrus,

Kimmer O'Neill, Renee Edge, Christina Overstreet, Bonnie and Gene Parrish, Ulrich Bernier,

Reginald Coler, and Eugene Gerberg. Finally, thanks to all the persons involved with the US

Navy's Duty-Under-Instruction (DUINS) program that provided the opportunity to attend

graduate school while serving as a Naval preventive medicine officer.









TABLE OF CONTENTS

page

A CK N O W LED G M EN T S ................................................................. ........... ............. .....

L IST O F T A B L E S ................................................................................................ 8

LIST OF FIGURES .................................. .. ..... ..... ................. .9

L IST O F A B B R E V IA T IO N S ............................ ................................................... ...................10

A B S T R A C T ................................ ............................................................ 12

CHAPTER

1 INTRODUCTION AND LITERATURE REVIEW ................................... .................14

Introdu action ................... .......................................................... ................. 14
T ru e F lies .................. ............... ................................................................ 15
M edically and Economically Important Flies...................................................................... 16
H o u se F ly ................... ...................1...................6..........
N om en clatu re ................................................................17
Origin and D distribution ............................................... .. ..... ................. 19
Classification and Taxonom y .......................................................... ............... 19
Biology and Ecology ..................................... .............. ......... 20
L ife -cy c le ......................................................................... 2 1
Breeding substrates .................................. ... .. .................... 26
N nutrition and diet ............................................ .. .. ............. ......... 27
B a cteria a s F o o d ................................................................ ...............................2 8
M icrobiology ............................................. .. ........... ......... 30
Microorganism persistence, replication and genetic transfer...............................31
Development of antibiotic-resistance in house fly gut...........................................33
Insect M ovem ent ......................... ......................... ............................35
Mark-Release-Recapture Studies and Techniques .................................. ...............36
Flight and D ispersal ....................................................... .......... .. ............ 37
Mark and Release Techniques.................. ................................ 39
Population Dynamics: Monitoring House Fly Populations...........................................41
Passive m monitoring m ethods.......................................................... ............... 41
A ctive m monitoring m ethods............... ..................................................................44
House Fly Economical Impacts and Disease Outbreaks ..............................................47
H ou se F ly M anagem ent.......................................................................... ................... 48
C u ltu ral control ................................................................4 8
M mechanical control .................. ............................. ........ .. ........ .... 49
Biological control ......................................... .................... .... ...... 51
C h em ical control ...............................................................53
Integrated P est M anagem ent ........................................ ............................................54
U usefulness of H ou se F lies .................................................................. .... ...................55









Enterobacteriaceae ............... .............................. ............................ 57
E sch erich ia co ..............................................................................5 8
P athogenic E scherichia coli ....................................................................... ......................6 1
E sch erich ia co li 0 157 :H 7 ............... .................................... ........ ................ .... ........... 63
Escherichia coli 0 157:H 7 and Cattle................................................ ......... ............... 63
Escherichia coli 0157:H7 Outbreaks.............................. ......... ...............65
Escherichia coli 0 157:H7 Pathogenicity ............................................. ............... 67
Escherichia coli 0157:H7 Prevalence and Persistence.......................... ...............68
Escherichia coli 0157:H7 Detection, Isolation and Identification ..............................69
Escherichia coli 0157:H7 and DNA-based Isolation Techniques..............................76
S u m m ary ................... ...................7...................7..........

2 H O U SE FL Y D ISPE R SA L .......................................................................... ....................78

In tro d u ctio n ................... ...................7...................8..........
M materials and M ethods ........................ .. ........................ .. .... ........ ........ 83
Laboratory Facilities and Rearing ............................................................................83
D description of Study A rea .......................................................................... ............... 84
H house Fly Collection and Rearing ............................................................................ 88
Transport of A dult Flies to the Field ................................. ........................................... 92
Marking, Releasing and Recapturing Adult House Flies .........................................92
Effects of Fluorescent Dust on House Fly Adults.........................................................96
W weather ......................................................................................... .... 98
Statistical A nalysis................................................... 99
R e su lts ................... ...................9...................9..........
D iscu ssion ...... .........................................................102

3 ESCHERICHIA COLI 0157:H7 PREVALENCE ......... .....................131

In tro du ctio n ........................131.....................................
M materials and M methods ...............................................................134
D N A Q u an tificatio n ................................................................................................ 15 1
16 S rD N A P C R A naly sis ........................................................................................ 152
Statistical A analysis .............................................. 152
R e su lts ........... .. ........ ......... ............................................................................................. 1 5 4
Fly Monitoring...................... ....... ..... ..................154
Enumeration of Aerobic Bacteria and Escherichia coli 0157:H7 .......................... 155
Escherichia coli 0157:H7 Prevalence by Direct Culture.............................156
Escherichia coli 0157:H7 Prevalence by Polymerase Chain Reaction ...... ......157
D iscu ssion ...... .........................................................158

4 OVERALL CONCLUSIONS ................. ...... ......... .........179

C onclusions.....................................................................179
B a c k g ro u n d .............................................................................................................. 1 7 9
C o n c lu sio n s .............................................................................1 8 0
Future Research .............. .. ...... ....................................................................... 181



6









Sum m ary ................... ................... ............................81

LIST O F R EFER EN CE S ............................................................................. ..........................183

B IO G R A PH ICA L SK ETCH ............................................................................. ....................215



















































7









LIST OF TABLES


Table page

2-1 Mean and maximum distances flown per week and year by marked house flies
released at a dairy in north central Florida ....... ....... .......... ................. ............... 125

2-2 Trap distances (km) from the release site, the total numbers of marked house flies
captured per alsynite sticky trap, and the cumulative percentage of marked flies.. ........126

2-3 Weekly recapture rate of marked house flies on alsynite sticky traps ..........................128

2-4 Total number of house flies and marked house flies captured on alsynite sticky traps
following release at a dairy farm in north central Florida. .........................................129

2-5 House fly release week and recapture rate and associated weekly weather data.............130

3-1 Dates and house fly monitoring methods used at two Florida dairies...........................173

3-2 Primer nucleotide sequences used to amplify target genes in PCR assay .....................174

3-3 Enumeration of aerobic bacteria (CFU/g) using Petrifilm Aerobic Plate Count plates
inoculated with 1 ml of the unenriched sample. ................................... ............... 175

3-4 Mean enumeration, by site and by sample type, of aerobic bacteria (CFU/g) using
Petrifilm A erobic Plate Count plates.. ......................... ............................................. 176

3-5 Prevalence (%) and number ofE. coli 0157:H7 CHROMAgar-positive samples and
number of CT-SM AC and PCR-positive samples ............... ............ .....................177









LIST OF FIGURES


Figure page

2-1 Alsynite trap (Olson Products Inc., Medina, OH) placed at dairies and used to
recapture on-dairy and dispersing house flies....................................... ............... 122

2-2 Alsynite trap (Olson Products Inc., Medina, OH) locations and distance (km) from
release p point. ........................................................................... 12 3

2-3 Examples of dairy-collected-collected house flies following excessive treatment with
two dusts to determine 24 h mortality effects. .............................. ... ............124

3-1 Scudder grid (45 x 45 cm) used to assess house fly populations on dairy farms. .........169

3-2 Spot cards at Dairy A's milk barn and at Dairy B. .................................. .................170

3-3 Spot and sticky cards. ....................... ........................ .. .... ........ ......... 171

3-4 Sampling methods for each type of collected sample................................................172











ARS

BAMM

BM


BLAST


BMBL

CDC

CMAVE


CT-SMAC




DOC

EHS

FDA

FMRU

GHFD



HHS

IMS

IOWH

ITIS

MMWR

NCBI

NCDC


LIST OF ABBREVIATIONS

Agricultural Research Service. Scientific research agency of the USDA

FDA's Bacteriological Analytical Manual

Background microorganisms. Refers to competing microorganisms that
interfere with microbiological isolation of target organism

Basic Local Alignment Search Tool. NCBI web-based tool used to find
similar regions between nucleotide or protein sequences

Biosafety in Microbiological and Biomedical Laboratories

Centers for Disease Control and Prevention. Agency of HHS

Center for Medical, Agricultural, and Veterinary Entomology, located in
Gainesville, Florida. Research center of the USDA-ARS

Sorbitol-MacConkey agar supplemented with cefixime (15 [tg/l) and
potassium tellurite (1.25 [tg/l). Selective agar used to isolate Escherichia
coli 0157:H7 from samples. Addition of antibiotics increases specificity
over that of conventional SMAC

Department of Commerce

UF's Department of Environmental Health and Safety

United States Food and Drug Administration

Flies and Mosquitoes Research Unit. Research unit at CMAVE

Gainesville (larval) house fly diet. Standard rearing medium at the USDA-
ARS-CMAVE for immature house flies. Consists of 50% wheat bran, 30%
alfalfa meal, and 20% cracked corn (Hogsette 1992)

United States Department of Health and Human Services

Immunomagnetic separation

Institute for One World Health

Integrated Taxonomic Information System

CDC's Morbidity and Mortality Weekly Report

NLM's National Center for Biotechnology Information

NESDIS's National Climatic Data Center









NESDIS NOAA's National Environmental Satellite, Data, and Information Service

NIH National Institutes of Health

NLM NIH's National Library of Medicine

NOAA DOC's National Oceanic and Atmospheric Administration

OHS CDC's Office of Health and Safety

SMAC Sorbitol-MacConkey agar. Selective agar used to isolate Escherichia coli
0157:H7 from samples

UF University of Florida

USDA United States Department of Agriculture

USN United States Navy

WHO World Health Organization









Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy

MUSCA DOMESTIC L. (DIPTERA: MUSCIDAE) DISPERSAL FROM AND ESCHERICHIA
COLI 0157:H7 PREVALENCE ON DAIRIES IN NORTH-CENTRAL FLORIDA


By

Roxanne Grace Burrus

August 2010

Chair: Jerome A. Hogsette
Cochair: Phillip E. Kaufman
Major: Entomology and Nematology

House fly, Musca domestic L., dispersal up to 3 km from a dairy release site into a nearby

town was documented in this study. Dispersal occurred by both direct flight across multiple,

mixed habitat types including open fields with interspersed tree copses, and by corridors and

edges provided by local roads. Marked flies were collected at an adjacent dairy, and at most traps

that were set along two well-travelled roads connecting the dairies and the town. Additionally,

one marked fly was captured on a trap that was placed outside of a restaurant in town 3 km from

the release site. In total, 0.67% (250) of marked house flies were recaptured from a total release

of 37,2000 marked flies over 11 wk.

Escherichia coli 0157:H7 was isolated from two dairies in north-central Florida using

immunomagnetic separation followed by direct culture plating onto CHROMAgar and sorbitol

MacConkey agar supplemented with potassium tellurite and cefixime (CT-SMAC) selective

media. Presumptive identification of E. coli 0157:H7 was confirmed by polymerase chain

reaction (PCR) using the fliCH7 and rfbEo157 gene amplification.

Dairy samples that were tested included pools of house flies, spilled grain, and fresh dairy

cattle manure. Forty-two percent (24/57) of samples were positive by direct-culture using both









CHROMAgar and CT-SMAC agar, with 11 positive samples from CHROMAgar and 13 positive

samples from CT-SMAC agar. The 24 positive samples were submitted to PCR; of those 24

samples, 14 (58%) were confirmed by PCR. Two of the PCR-confirmed samples were false-

negatives on CHROMAgar media, but presumptive positive on CT-SMAC media, indicating the

importance of analyzing samples by more than one method, and demonstrating the sensitivity of

PCR.

Direct culture prevalence rates from CHROMAgar were 14.0% (8/57) for house flies,

5.3% (3/57) for grain, and 0% (0/57) for manure. The rate of CHROMAgar isolation ofE. coli

0157:H7 from house flies was 2.6 times greater than from grain. The PCR confirmation rates

were 67% (8/12) for house flies, 56% (5/9) for grain and 33% (1/3) for manure. These data

suggest that detection ofE. coli 0157:H7 on dairies might be more accurately determined by

testing house flies instead of grain or manure samples, regardless of which isolation method is

utilized.

Flies are easy to collect and process, and because they disperse into urban areas, they

provide valuable information regarding a mobile element for pathogen transmission that is

lacking in grain and manure samples. House flies should be an important consideration in the

design of a pathogen monitoring program on dairies.









CHAPTER 1
INTRODUCTION AND LITERATURE REVIEW

Introduction

With projected increases in global and local temperatures (West 1951, Borror et al. 1989,

Gullan and Cranston 2000, Goulson et al. 2005, Meerburg et al. 2007), there will likely be a

concomitant increase in insect vector populations, such as filth flies (Gratz 1999). This may

potentially cause an increase in diarrheal diseases transmission by enteric bacteria such as

Escherichia coli Castellani and Chalmers, .\lngell/ spp., and Salmonella spp., due to mechanical

transmission of pathogenic organisms by filth flies (Greenberg 1971, Greenberg 1973).

Filth fly life-cycle developmental times for each developmental stage decrease in

duration with increased temperatures; as a result, the number of generations per year increases,

and in temperate areas, can result in the establishment of year-round fly populations. Higher-

than-normal, year-round filth fly populations, living in close proximity to human populations,

carrying viable antibiotic-resistant bacteria in their digestive tracts, and are themselves resistant

to multiple pesticides (Kaufman et al. 2001), present a tremendous potential for significant

increase in human disease. Petridis et al. (2006) documented E. coli 0157:H7 genetic transfer of

antibiotic resistance within the house fly, Musca domestic Linnaeus, gut. Additionally, house

flies can travel as far as 8 km (13 mi) from their breeding sites (Bishopp and Laake 1921, West

1951, Quarterman et al. 1954, Sacca 1964, Stein 1986, Milio et al. 1988). This increases the

potential for introduction of diseases across species, particularly from dairies with large house fly

populations (Kaufman et al. 2005), to nearby human population centers.

Due to anticipated expansion of existing urban areas and the simultaneous expansion of

animal facilities, large animal-rearing facilities with concomitant filth fly populations will be

increasingly in closer proximity to large human populations. The close proximity becomes a









significant human health threat when the animal and human populations are within the filth fly's

normal flight distance, due to the increased opportunities available to the fly. This is of special

concern in urban areas with hospitals producing biohazardous human waste, and in areas where

both farms and urban centers generate antibiotic-resistant bacterial pathogens which can then be

transferred between sites. The goal of this research is to determine the public health threat posed

by house fly, transmission of enterohemorrhagic Escherichia coli, across the rural-urban

interface between rural dairy farms and nearby urban residential areas.

True Flies

The house fly is a true fly. All true flies are in the Order Diptera, (Latin, di "two" +ptera

"wings"), with hind wings that have been reduced to clubbed halteres. The halteres serve as

balance-sensory organs and project from the mesothorax at the site where hind wings would

normally be located. Thus, true flies have only one pair of true wings (Borror et al. 1989).

Possession of only one pair of wings, accompanied by the presence of halteres instead of the

typical insectan characteristic of two pairs of wings, is a characteristic unique to Diptera, and

serves as an easily-observed visual diagnostic for differentiation of adult flies from most other

adult insects. There are some exceptions to the one-pair of wings characteristic: some insects in

other orders, e.g., some mayflies (Ephemeroptera), have only one pair of wings, but lack halteres

(Borror et al. 1989). A more notable exception are male scale (Coccoideae) insects. Like Diptera,

male scale insects possess halteres; however, scale halteres are bristled hooks instead of clubbed

processes typical of true flies. Additionally, male scale insects lack mouthparts, and typically

have one (rarely two) style-like process projecting from the abdominal tip (Borror et al. 1989).

There are 108 families of true flies (Olsen 1998), with an estimated 100,000 (Smith 1986)

to 120,000 (Borror et al. 1989) identified species in the world. Identification of new species

during the past20 years has increased that number to more than 150,000 fly species (Thompson









2009). Diptera are one of the largest orders of insects. Large, diverse populations are typical in

most locations (Borror et al. 1989) with more than 20,000 species of Diptera in the Nearctic

Region alone (Mullen and Durden 2002).

Medically and Economically Important Flies

Of the more than 150,000 true fly species (Thompson 2009), 350 (Greenberg 1971) are of

medical, veterinary, and/or economic importance to humans, either because they transmit the

pathogens that cause disease, or because they interfere with animal-rearing and crop production.

Many of these flies are synanthropic, whereby they exhibit such strong associations with humans

and domestic livestock that they are typically found living only with humans and in human

environments (Borror et al. 1989). Fewer than 3.5% of flies representing fewer than 350 species

in 29 families are associated with pathogen transmission. (Greenberg 1971). One species, M.

domestic (Diptera: Muscidae), is so synanthropic that it is commonly referred to as the house

fly (Howard 1900, Hewitt 1910, Hatch Jr. 1911, Parkes 1911, Mullen and Durden 2002).

House Fly

Flies have had a major impact during wars, due to transmission of enteric pathogens that

cause diarrhea, dysentery and typhoid fever. Some war-related examples follow. In 1898, the

role of flies in pathogen transmission was discovered during the Spanish-American War (ANON

1940), as more soldiers died from typhoid fever, which is transmitted by house flies, than from

battle injuries (Cirillo 2006). This discovery resulted in the application of the name "typhoid fly"

to M domestic and the publication of a 2-volume report by a commission of medical officers

chaired by Walter Reed (ANON 1940). Combatants in the Anglo-Boer War, also waged in the

late 1890's, suffered more deaths from typhoid fever than from battle injuries (Cirillo 2006).

Subsequently, public health efforts were waged in the United States against the typhoid fly.









Walter Reed and his commission determined that M. domestic was predominantly responsible

for the transmission of typhoid fever (ANON 1940, Cirillo 2006).

During World War II, in 1942, United States Army camps suffered a "massive epidemic

of bacillary dysentery." It was determined by serological and biochemical cultures that the same

species of bacteria was present in the flies and in more than 91% of the infected soldiers.

Importantly, no flies had been observed at the bivouac site prior to the Army's arrival. However,

cases of dysentery and numbers of flies increased rapidly. After fly control measures were taken,

the number of flies and the incidence of dysentery both decreased. Although no mortality

occurred during this epidemic, 22% of the soldiers in the Division tested positive for the

pathogenic organism. Infected persons were incapacitated for two to seven days.

Changes in technology led to reduced breeding sites for house flies that in turn led to

reduced numbers of flies and fewer concomitant cases of diseases in which flies were implicated.

For example, Graham-Smith (1939), as cited in ANON (1940) attributed the decrease in summer

diarrhea of infants to the increased use of automobiles with concomitant decreased use of horse-

drawn carriages.

Nomenclature

Nomenclature of flies has varied over time, among countries, and between professional

organizations. For example, in the United States, current convention for entomological

terminology (Borror et al. 1989) in the largest North American entomological organization, the

Entomological Society of America (ESA), specifies that common names for true flies include the

word "fly" as a separate word: e.g., house fly, stable fly, horn fly, deer fly, horse fly, sand fly,

etc. Correspondingly, this ESA convention specifies that insects in other orders, which are not

true flies, append the suffix "-fly" to the common name root word: e.g., sawfly, stonefly,

caddisfly, dragonfly, damselfly, etc. In contrast, older North American literature (before 1970),









often cites "the housefly," as one word (Matthysse 1945, Olson and Dahms 1945, Wilkes et al.

1948, West 1951, Floyd and Cook 1954, Sacca and Rivosecchi 1958, Sacca 1958, Greenberg

1959a).

Disparity of common names was a very big problem in much of the early literature, and

can make historical research difficult. Two of the largest and most prominent entomological

organizations in the United States formed standing committees just after the turn of the 20th

century to resolve this matter: the American Association of Economic Entomology (AAEE) in

1903, and the American Entomological Society (predecessor to the current ESA) in 1907. The

AAEE published the first list of common insect names in 1908, in which it formally established

the use of "house fly," vs. "housefly," as the common name forM domestic. In 1940, the

AAEE and ESA insect nomenclature committees co-published this list of authorized common

insect names as a joint committee. In 1953, AAEE was absorbed into ESA and the resulting ESA

has continued publication of the list to the present. The list was originally published as a book

(Stoetzel 1989, Bosik 1977), but is now available only online as database (ESA 2009). A more

detailed description of the history of insect common name nomenclature is available for the

interested reader (Chapin 1989).

Unfortunately, while adherence to the common names in this database is required for

publication in ESA publications, many other peer-reviewed publications do not comply with this

attempt to unify nomenclature. This is particularly true in medical publications, but is also true

for some entomological journals that do not fall under ESA oversight. However, continued

emphasis of standardized and unified common name nomenclature does seem to have succeeded

in reducing the number of publications using non-standardized nomenclature; overall usage of









insect common names is fairly well established throughout most of the ESA peer-reviewed

literature in the United States.

In non-ESA publications worldwide, "housefly" is still in use today (Zarrin et al. 2007,

Malik et al. 2007, Black IV and Krafsur 2008, Cafarchia et al. 2009). Additionally, the reader

will find multiple common names used for a particular insect. One will find that the same

common name can refer to many different insects, depending on local nicknames found in

different geographical locations. There might be, as in the case of the house fly, multiple sub-

species of the insect that are so closely related they share not only their common name, but

Linnaean classification as well. For example, Musca domestic vicina Macquart, sometimes

referred to as M vicina, is the oriental house fly; however, it is often reported in the literature

simply as "house fly," thus confusing it with M domestic.

Origin and Distribution

The house fly is a cosmopolitan pest that is strongly associated worldwide with human

habitation, especially when domestic animals such as livestock, horses and poultry are present

(West 1951). For all practical purposes, wherever humans live, the house fly also resides. The

house fly is excluded from arctic regions and higher altitudes due to extended cold temperatures

(West 1951).

Classification and Taxonomy

Within the Order Diptera, the house fly is classified by Borror et al. (1989) as Suborder

Brachycera, Infraorder Muscomorpha (Cyclorrhapha), Division Schizophora, Section

Calyptratae, Superfamily Muscoidea, Family Muscidae, Genus Musca, and species domestic

Linnaeus 1758. Each of these classification levels reflects some morphological trait that is

characteristic for M domestic.









The Suborder Brachycera contains the short-homed flies (Borror et al. 1989), so named

because they typically have short antennae with five or fewer segments. In contrast, flies the

suborder Nematocera typically have long narrow antennae with 13 or more segments (Borror et

al. 1989). Placement in Infraorder Muscomorpha (Cyclorrhapha) is based partially on the method

of adult emergence from the puparium. The Muscomorpha includes the more advanced and

specialized (higher) Diptera, which emerge through two slits that together appear "T-shaped" on

the anterior portion of the relatively stout, oval puparium (Jones 1977). Muscomorpha open these

slits by alternately expanding and contracting a specialized inflatable sac (ptilinum) that is

present on the head of emerging cyclorrhaphous flies. The pressure of the expanded ptilinum

against the puparium causes the slits, only a few cells thick, to part so that a roughly circle-

shaped opening is formed from which the fly emerges. The ptilinum retracts into the head shortly

after emergence, but a seam (frontal suture) that is described both as being circular (Jones 1977),

or as being shaped like a horse-shoe (Hogsette, personal communication), remains visible in the

adult fly (Jones 1977). The presence of a frontal suture in adult flies is responsible for

classification into the Division Schizophora. In contrast, Aschiza adults typically lose their

ptilinum and lack a frontal suture. Schizophora are divided into two sections (Calyptratae and

Noncalyptratae) based on possession or lack of a pair of calypters (squamae, alulae) and a

thoracic transverse suture (West 1951). Calyptrate flies possess well-developed calypters that are

large enough to conceal the halteres and they have an easily-observed transverse suture.

Biology and Ecology

The literature is replete with bionomic information concerning the house fly and closely

related filth flies. Excellent monographs on M. domestic are available (Hewitt 1914, West

1951). A comprehensive monograph on the synanthropic black blow fly, Phormia regina

Meigan, provides excellent information regarding the physiological aspects of eating that are









also illustrative forM. domestic (Delthier 1976). Greenberg (1971, 1973) wrote a thorough two-

volume series that specifically addresses flies and pathogen transmission. Although much of the

early literature is dated, these monographs provide tremendously useful information that is not

easily available in the current literature. The wealth of information provided by these authors is

tremendous, and should not be neglected by anyone interested in studying the house fly or

related muscids.

Life-cycle

House flies have a holometabolous life-cycle (Borror et al. 1989), which consists of four

stages: egg, larva, pupa, and adult. Development of house flies at each life stage is temperature-

dependent, so that the entire life-cycle can be completed in as few as 8 d in the summer (West

1951). However, the typical life-cycle for development from egg to adult in temperate zones

ranges from 10-14 d. First stage larvae hatch from the eggs within 8-12 h after oviposition. The

three larval stages are typically completed as follows: first instar in 20 h 4 d, second instar in

24 h to several days, and third instar in 3-9 d. The pupal stage lasts an average of 5 d, although

adverse conditions may extend this to several weeks. The adults live several weeks, with females

generally living longer than males. During summer, adults may live 2-3 wk while cooler

temperatures during spring and fall may contribute to adult longevity for up to 3 mo (West

1951).

House fly eggs have a glossy white (West 1951) or creamy color (Mullen and Durden

2002), are approximately 1 mm in length (West 1951), but can range in size from 0.8-2.0 mm

(Mullen and Durden), and are approximately 0.25 mm in width (West 1951). House fly eggs

exhibit a slight concave indentation on the dorsal side that is due to the presence of two

longitudinal ridges (ribs) that taper towards each other anteriorly (West 1951, Mullen and

Durden 2002) to form hatching pleats (Mullen and Durden 2002). Gravid females disperse their









eggs methodically throughout available breeding substrates, depositing them either individually

or in masses (West 1951). The females walk over and crawl carefully through the breeding

substrate, seeking cracks and crevices into which they extend their ovipositor to dispense ova.

The ova are laid so that they "rest on their broader posterior ends" (West 1951). Females oviposit

4-8 d after mating, and may take up to 24 h to deposit each batch of 100-150 eggs. A total

lifespan deposition of 4-6 batches of eggs can occur when flies are permitted to oviposit every

two weeks (West 1951). Flies that are maintained in colony are induced to lay eggs every other

day after they are five days of age (Hogsette, personal communication).

House fly larvae are also called maggots (Moon 2002). House fly larval length increases

approximately 25 percent during each of three molts and body weight can increase 54-fold in 4 d

(West 1951). Third instars are approximately 12 mm (West 1951) to 15 mm (Moon and Meyer

1985) in length. House fly larvae possess two fused (Roback 1944) mouth hooks in their reduced

(Moon 2002), unsclerotized head (Borror et al. 1989) that is located at the anterior, pointed end

of a tapered body. The head lacks eyes but possesses small papillae-like antennae (Moon 2002).

The larvae are creamy white in color (Moon 2002). The mouth hooks move in a vertical plane

(Borror et al. 1989), providing physical evidence that these larvae are non-predaceous, because

their mouth hooks do not permit grasping. Instead, these mouth hooks permit the larvae to pull

themselves through the media in which they live. House fly larvae are primarily sarcophagous,

i.e., they feed on rotting organic matter (West 1951) (Gr. sapros, rotten; phagein, to eat). House

flies are also reported to be coprophagous, i.e., they feed on dung (Hammer 1941) (Gr. kopros,

dung). The paired mouth hooks can be retracted into the oral cavity or extended out, and are used

to help the larva move through food substrates (Moon 2002). House fly larvae are legless (Borror

et al. 1989). Each of the eight abdominal segments has a transverse row of spines on the ventral









surface (creeping welts) that are used to facilitate movement (Moon 2002). House fly larvae

possess a pair of lateral spiracles on the prothorax of second and third instars and a pair of

spiracles at the last abdominal segment of all instars (Moon 2002).

The posterior (caudal) spiracles and associated structures are useful for aging and

identifying muscid immatures, especially in third stage larvae (Moon 2002). Posterior spiracles

are located on a spiracular plate which is encircled either completely or partially by a sclerotized

peritreme. Developmental age can be determined to instar by counting the number of spiracular

slits. First and second house fly instars possess two spiracular slits inside the spiracular plate,

while third instars possess three slits.

House fly larvae can be identified to species by examining the shape and orientation of

the spiracles, the location of the molt scar, and the peritreme shape. House fly larvae have

sinuous spiracles partially surrounding a large molt scar. The peritreme is completely sclerotized

and encircles the spiracular area, with the peritreme flattened along the interior edge and the molt

scar located mid-way along the interior flattened edge of the peritreme. Closely related muscid

larvae such as the stable fly, Stomoxys calcitrans (L.), with similar body shapes and sizes can be

differentiated by s-shaped spiracular slits surrounding a small molt scar that is located in the

center of a spiracular plate that lacks a flattened interior edge (Moon 2002).

Mature 3rd instars migrate from the deeper moist, fermenting parts of the larval medium

up into drier crusty top-layers (Greenberg 1959d). Once there, they wander for approximately 24

hours, during which time they cease feeding in preparation for pupation so that the prepupal gut

is empty and contracted (Greenberg 1959d).

The third instar's integument becomes hardened during the process of pupariation, to

form a puparium, in which the fly will complete its development (Moon 2002). House fly pupae









are accurately called puparia (West 1951), to reflect the enclosure of the exarate coarctatee) pupa

within the sclerotized last larval cuticle, in contrast to larval metamorphosis within a specially

constructed pupal structure that is typical for insects other orders such as Lepidoptera and

Coleoptera (Gullan and Cranston 2000). The puparium is red to brown in color and barrel-shaped

(Moon and Meyer 1985) and typically 6.3 mm in length (West 1951). Pupariation takes at least 6

h, with sclerotization gradually darkening the integument to its final dark brown color during this

process (West 1951).

The pupal stage typically lasts 5 d (West 1951), but can range from 3-10 d (Moon and

Meyer 1985), depending on temperature (West 1951, Moon and Meyer 1985). During adverse

conditions, the pupal stage can last several weeks, and pupae may be able to hibernate (West

1951). Pupae are usually located in a cooler (West 1951) and drier (Moon and Meyer 1985)

location than feeding larvae. Third instar nonfeeding prepupae disperse to find suitable locations

for pupal development. Large pupal aggregations numbering in the thousands can be found close

to the medium surface and in surrounding soil (Moon and Meyer 1985).

Pupal development lasting 4 d at 35 C (Greenberg 1959b) occurs with daily

physiological changes. Within the first day, the pupa molts, and is encased in a transparent

molting membrane with the head uneverted. On the second day, the head everts, and the

compound eyes are amber. On the third day, the eyes are orange or brown and bristles exhibit

slight pigmentation. On the fourth day, the fly ecloses (Greenberg 1959b).

House fly adults are small to medium sized flies approximately 6mm in length (West

1951). However, they may range in size from 3-10 mm (Byrd and Castner 2009) or from 4-12

mm (Moon 2002). Numbers of adults and adult size are reduced by inadequate larval nutrition

(West 1951) or by high larval population density (Sokal and Sullivan 1963, Black and Krafsur









1985). Density-induced adult size changes can influence mating success (Baldwin and Bryant

1981).

The house fly life-cycle length can vary due to one or a combination of multiple factors

such as: temperature (Hewitt 1910), relative humidity (Hewitt 1910), and light (Greenberg

1959e). Additional factors that can influence developmental times include geographical region

(Black and Krafsur 1968); season (LaBreque et al. 1972); breeding substrate composition (Skoda

et al. 1993), pH, and moisture content (Evans 1916); population density (Haupt and Busvine

1968, Taylor and Sokal 1977); and nutrition (West 1951).

House flies begin their life-cycle shortly after adult emergence. Males and females

emerge in approximately equal proportions when adequate nutrition is available (West 1951).

When insufficient nourishment is available, significantly smaller males develop and these greatly

outnumber the females (West 1951).

The time required for sexual maturity in newly emerged adult house flies was reported as

1 d by Riemann et al. (1967). Murvosh et al. (1964) found that males are sexually mature at 16 h

and females at 24 h. A great deal of research has been conducted to determine the preoviposition

period of house flies, for the purpose of concentrating control efforts against adults before

oviposition occurs (Hutchison 1916). Hutchison (1916) determined that preoviposition periods

range from 2.5 23 d, and are greatly influenced by temperature. His data imply that

preoviposition periods also may be influenced by humidity and adult diet. Bishopp et al. (1915)

also studied preoviposition periods, and they observed a seasonal / temperature influence on the

preoviposition period, i.e. the period between emergence and oviposition, during which the fly

becomes sexually mature and copulation takes place (Mellor 1919). Bishopp et al. (1915)









observed preoviposition periods of 4 d in summer vs. more than 10 d in spring and fall, and

Hewitt (1910) observed long preoviposition periods of up to 14 d.

Several days elapse between sexual maturity and copulation, so that females typically

mate when 3-4 d old (West 1951). Adult female house flies are typically monogamous, with only

a small percentage mating more than once, and none more than a few times. Monogamy appears

to occur due to a threshold limit for the receipt of seminal fluid (not of sperm) which can be

reached by interrupted matings. Matings with castrated males can also result in loss of

receptivity. Females may become sexually receptive again after 20 consecutive days of

oviposition. Transfer of sperm is essentially completed within 10 minutes, although flies remain

coupled for approximately 1 h. Even those adult female flies that mate more than once restrict

their copulations to only a few times (Riemann et al. 1967). Flies copulate with the smaller male

positioned directly above and clasping the female, with both insects facing the same direction,

and with their abdominal tips connected (West 1951).

Breeding substrates

House flies are cosmopolitan opportunists, and are associated with a wide variety of

decomposing organic materials (Lole 2005). Availability of a natural substrate is the most

important factor for house fly selection at any particular location and in a specific moment.

Natural substrates consist of a tremendous variety of materials, including manure from cattle,

horses, sheep, dogs, and humans; spilled grains; compost; garbage and landfills (Lole 2005).

House flies are ubiquitously abundant on cattle farms, due to the availability of preferred

breeding sites such as decaying grain-based feed and manure (Skoda et al. 1993).

Larvae spend their entire developmental life-cycle within the substrate selected by the

gravid females. In general, house fly larval abundance in substrates reflects localized abundance

of that particular substrate. For example, prior to the development of the automobile, enormous









house fly populations were observed in horse manure that was greatly abundant in urban streets.

Post-automobile, we find very few references to house flies breeding in urban horse manure.

Currently, we find that house fly problems at animal-rearing facilities, rather than in urban

streets, dominate the literature. Current fly problems coincide with increased numbers of ever-

enlarging commercial facilities, which struggle to manage the copious amounts of fecal waste

generated by the intensively-reared animals.

Nutrition and diet

Survival rate at each developmental stage, pupal weight, and adult fecundity are

important measures of the nutritional status of reared flies. Larval populations that do not receive

adequate nutrition usually produce smaller flies with higher proportions of males. House fly

larvae are successfully maintained in the laboratory on a variety of diets. After nutritional

provision, important factors to consider in selecting a rearing medium are ease-of-use, expense,

and inhibition of molds and fungi.

Sukhapanth et al. (1961) compared fly development in a synthetic diet ("100 g rice bran

and husk, 350 g dry, low-fat powdered milk, 75 icing sugar, 15 g Baker's yeast and 200 ml of

2% KOH in normal saline solution"), a natural diet ("fresh cow meat"), and a combination of the

two. House fly egg production and survivorship was highest at all developmental stages on the

combined synthetic/meat diet and lowest at all stages on the synthetic diet. Sukhapanth et al.

(1961) did not describe pupal weights, or adult sizes, so that comparison to diets used by other

researchers cannot be made without repeating Sukhapanth's work.

Hogsette (1992) created the Gainesville house fly diet (GHFD) containing 30% alfalfa,

50% wheat bran, and 20% corn meal to take advantage of year-round locally-available feed

components. Mean larval weight (15 g) and adult eclosion rates from the GHFD diet did not

differ significantly from that of the Chemical Specialties Manufacturers' Association larval









medium (CSMA) (Greenberg 1959b). CSMA is composed of 33% wheat bran, 26.7% alfalfa

meal, and 40.0% brewer's grain (Greenberg 1959b). Additional benefits that Hogsette (1992)

described for use of GHFD over CSMA were decreased costs, reduced delivery times, decreased

storage time, increased feed quality, and suitability for stable fly, Stomoxys calcitrans L. larvae

by addition of peanut hulls to the GHFD in equal volumes.

Bacteria as Food

The nutritional needs of house fly larvae are fulfilled by consumption of micro-organisms

and other substances located within the decomposing organic matter that also serves as a

breeding substrate. Some natural components of substrates used for larval development ofM.

domestic are microorganisms, such as bacteria, viruses, and parasites. Postulation that house

flies eat bacteria, and not just the decomposing substrate materials used for larval development,

led to experiments to rear M domestic in various synthetic media. Schmidtmann and Martin

(1992) determined that house fly larvae depend either on the bacteria or bacterial metabolic

products for essential nutrients. In some studies, agar-based systems with known microbial

organisms have been used to rear house fly larvae (Schmidtmann and Martin 1992, Watson et al.

1993, Lysyk et al. 1999) and larvae were observed during development (Perotti and Lysyk

2003).

Bacterial communities of decomposing organic substrates can change rapidly, due to a

series of complex biotic and abiotic interactions (Archer and Young 1988, Jiang et al. 2002).

Environmental conditions such as temperature and relative humidity affect several substrate

factors, including decomposition rate, available water content, pH, and availability of oxygen.

Bacterial communities can also be impacted by the metabolic processes of the competing

microorganisms (Jiang et al. 2002).









Investigation into microbial contributions towards house fly larval development has led

to experiments in which many specialized diets are used for rearing the larvae. These specialized

diets are narrowly defined by specific nutritional terminology such as axenicc," "gnotobiotic,"

"holidic," and "meridic." Because some researchers are describing the substrate as a rearing

medium rather than defining the nutritional composition of the medium, words such as "aseptic"

and "sterile" have alternatively been used. Usage of these words is not consistent among

researchers, and comparison of rearing methods must be carefully assessed by perusal of the

scientist's methods rather than by the word used to describe the larval rearing medium.

Some subtle, but important differences separate each of these words. Although axenicc"

and "aseptic" are frequently used synonymously in arthropod literature (Rodriguez 1966); both

terms refer to the rearing of target organisms with no other living organisms present, e.g., rearing

sterilized house fly in media containing only inert nutrients such as synthetic diets or nutrient

agars. The composition of inert components might or might not be specified. "Sterile" is

synonymous to axenic and aseptic, as seen in the rearing of house fly larvae on synthetic egg

yolk media and on blood agar plates that contained no other living organisms (Watson et al.

1993). "Gnotobiotic" refers to rearing of the target organism with only one other living organism

present, e.g.: a gnotobiotically reared house fly is an aseptic fly that was fed known quantities of

precisely defined foods, such as pure cultures of either Salmonella typhimurium or Proteus

mirabilis in known concentrations (Greenberg et al. 1970, Watson et al. 1993). Many studies that

are termed axenicc" would fall under the more specific term "gnotobiotic" due to provision of

bacteria species; thus, it seems that the term gnotobiotic has been generally supplanted by the

term axenic. Regardless of gnotobiotic status, axenic studies provide information about

nutritional responses (Rodriguez 1966) and have been used to determine the relationship of









bacteria and immature fly development by many researchers including Schmidtmann and Martin

(1992), Watson et al. (1993), and Lysyk et al. (1999).

The terms "holidic" and "meridic" were introduced by Dougherty (1959), as cited in

Rodriguez (1966), to differentiate between diets that contain only components with completely

known chemical structures (holidic) or diets that add components with unknown chemical

structures, such as agar, to a holidic base (meridic) (Rodriguez 1966).

Microbiology

Musca domestic hosts a large variety of bacterial fauna, both internally and externally

(Greenberg 1973). Due to the house fly's association with a wide variety of fecal and

decomposing organic materials, what can be considered natural fauna versus environmentally

obtained fauna is unclear. Despite this, Greenberg (1959c) postulated that Proteus vulgaris,

Proteus mirabilis, Aerobacter aerogens and Escherichia freundii might be considered the

"normal flora" of house flies, after he isolated these bacteria from both laboratory flies reared on

the CSMA diet and from natural populations of flies collected from horse manure. He further

concluded that these bacteria are probably the predominant species in CSMA

The advent of the microscope conferred the ability to study microorganisms, while

culture methods provided the ability to grow microorganisms. Used together, these two

technologies contributed greatly to the progress that has been made in isolating pathogens from

M. domestic and related flies. Since then, the house fly has been determined to be capable of

transmitting more than 100 microorganisms, including bacteria, parasites, viruses, and yeasts

(Greenberg 1973). Many of the microorganisms isolated from house flies are pathogenic to

humans.









Microorganism persistence, replication and genetic transfer

Extensive research has been conducted to examine the ability of microorganisms to

survive (persist) (Ledingham 1911; Graham-Smith 1912; Greenberg 1959b, 1959c; Sasaki et al.

2000; Zurek et al. 2001; Nayduch et al. 2005) and multiply (replicate) within the house fly's

alimentary canal and associated organs (Petridis et al. 2006, Macovei et al. 2008). The ultimate

goal is to determine the potential for biological (versus "merely" mechanical) transmission of

pathogenic agents by the house fly.

Microorganisms are consumed by larval house flies which depend on the pathogens or

their metabolic products for growth and development (Greenberg 1959b). Microorganism

survival from the larval to the adult fly stages could result in increased dissemination by the

house fly (Greenberg 1959b,d,e). Greenberg (1959c,d,e) observed large declines in the number

of bacteria in the prepupae and emergent adults. He attributed the decline in prepupae to a

cessation of feeding with continued excretion prior to pupation. This was confirmed by

observation that the house fly prepupal gut is empty and contracted in contrast to the fully

distended gut of actively-feeding larvae (Greenberg 1959d). The decline of bacterial counts in

newly emergent flies was attributed to the molting of foregut and hindgut during pupation

(Greenberg 1965). This was confirmed by counting the number of bacteria present in shed

puparia (105) versus the number of bacteria on the newly emerged flies (102). Greenberg (1965)

reported that the bacteria survived the pupal period within the shed foregut and hindgut portions

outside of the pupal fly, and were separated from the fly by a thin membrane. Therefore, adult

flies emerged from the puparium with relatively few bacteria. Those bacteria that did remain

within the fly were in the midgut, which is not shed during molting (Greenberg 1965).

Bacteria were reported to survive in the adult fly for up to 12 h on the surface, and up to

30 h in the gut and feces (Grubel et al. 1997). Sasaki et al. (2000) reported survival of









microorganisms for up to 3 d in the mouthparts and up to 4 d in the crop. The adult house fly

mouthparts consist of a proboscis, which has two fleshy labella that can be spread out, sponge-

like, over food surfaces, to implement a suctorial action for uptake of liquids and minute food

particles. Finally, the proboscis can be retracted into the head capsule, thus providing a micro-

habitat for bacteria that is relatively protected from desiccation and UV light (Kovacs et al.

1990). The proboscis also has one row of 5-6 three-cusped trifurcatedd) prestomal teeth on each

side of a central food channel (Macloskie 1880, Broce and Elzinga 1984, Sukontason et al.

2003). These prestomal teeth scrape food resources, and are important for enhanced pathogen

digestion (Kovacs et al. 190).

Microorganism survival within the house fly gut seems to differ for different

microorganisms. Nayduch et al. (2005) reported survival ofAeromonas caviae and Serratia

liquefaciens in the alimentary tract for up to 5 d. Nayduch et al. (2005) described production of a

bag-like peritrophic membrane structure around feces (fecal pellet). They observed that these

bacteria were located in the peritrophic membrane folds rather than in the peritrophic space, and

not within the ectoperitrophic space; indicating that they were not transported out with the feces

with the peritrophic membrane. They suggest that these bacteria are capable of evading

entrapment by the peritrophic membrane by some method (Nayduch et al. 2005). Viable

excretion of Yersiniapsuedotuberculosis was reported for up to 36 h with contamination of the

environment detected for up to 30 h after inoculation by Zurek et al. (2001). Griibel et al. (1997),

the first to detect viable Helicobacterpylori in house fly feces, reported survival of H. pylori for

up to 12 h on the exoskeleton, 30 h in the gut, and 30 h in the feces.

More than 100 spp. of bacteria have been isolated from adult house flies (Greenberg

1971), and from the adult's alimentary tract (Ahmad and Zurek 2006). The number of bacteria









on adults varies for different locations and bacteriological isolation methods. Sulaiman et al.

(2000) isolated nine spp. of bacteria, including the first reported isolation of Burkholderia

pseudomallei, from house flies in Kuala Lumpar, Malaysia.

Successful replication of bacteria within the digestive tract (Macovei and Zurek 2007)

and associated organs could play an important role in genetic transfer of virulence factors and

antibiotic-resistance from one bacterium to another, including transfer to different species,

because bacteria often exchange genetic material during replication. This is particularly true of

gram-negative enteric bacteria such as E. coli sp., which possess extra-chromosomal plasmids

(Johnson 2002).

Development of antibiotic-resistance in house fly gut

Insect guts provide an ideal environment for exchange of genes by plasmid transfer and

transconjugation between bacteria. The house fly has been implicated as a factor in the transfer

of antibiotic-resistance genes among bacteria (Macovei and Zurek 2006) because bacteria

replication, which is usually accompanied by genetic exchange, occurs in the house fly gut

(Petrides et al. 2006). Additionally, antibiotic-resistance was observed on organic pig farms

where antibiotics were not used, but fly numbers were large (Meerburg et al. 2007).

There is evidence that E. coli 0157:H7 exchanges genetic material such as virulence

factors and antibiotic-resistance by plasmid transfer (Perna et al. 2002) or phage-mediated

transduction (Johnson 2002). Plasmid transfer of antibiotic-resistance genes has led to disease

outbreaks that were difficult to control because of development of multidrug-resistant

Staphylococcus aureus (MRSA) (Perna et al. 2002). This bacterial exchange of genetic material

can occur within the house fly alimentary tract, which provides an environment conducive to the

evolution and emergence of new pathogenic bacterial strains (Petrides et al. 2006). Exchange of

antibiotic-resistance between infectious bacteria is responsible for the worldwide increase in









nosocomial (hospital-acquired) infections that are resistant to all known antibiotics (Weinstein

1998). The potential for bacterial exchange of genetic material within the house fly gut presents

potentially serious complications for any disease caused by pathogens that are disseminated by

house fly movement within a hospital environment. The potential for development of resistance

to multiple antibiotics is particularly troublesome if the flies have access to the pathogen's

natural reservoirs; for example, access to dairy cattle for transmission ofE. coli 0157:H7.

The potential for fly-borne bacteria dissemination coupled with bacterial exchange of

antibiotic-resistance is of concern in regards to the development of nosocomial infections

(Boulesteix et al. 2005). In hospitals, intensive-care units serve as the epicenter for these

infections (Weinstein 1998). While hospitals and animal-rearing facilities, such as dairies, both

report increased antibiotic-resistance due partially to over-use (drugs to humans for treatment of

bacterial infections and inclusion of drugs in animal feed for use as a growth stimulator),

dispersal of flying insects such as the house fly between hospitals and dairies also could

introduce new antibiotic-resistance genes in each site.

The potential for pathogen transmission, including antibiotic-resistance, plus the

possibility of economic losses due to infestations of synanthropic flies, has led to attempts to

reduce fly populations wherever flies aggregate or build to annoying levels. Insects that harm

human or animal health or damage valued resources are considered pests (Foster and Harris

1977). The importance of pest control for reduction of disease outbreaks is highlighted by reports

of paired reduction of fly populations and a corresponding decrease in the disease incidence

(Nash 1909, Lindsay and Scudder 1956, Cohen et al. 1991). This permits an estimated

quantification of the house fly's role in pathogen transmission. For example, Emerson (1999)

reports a 75% reduction of muscid flies corresponding to a reduction in the number of new cases









of trachoma and a 22-26% reduction of muscid flies corresponding to a reduction in the number

of new cases of diarrhea for villages in Gambia after fly control efforts reduced the numbers of

muscid flies by approximately 75%.

Insect Movement

Terrestrial insect movement can occur by walking or by flying. Flight is used to both

disperse and migrate (Angelo and Slansky 1984). Dispersal is typical of small, r-strategist insects

that produce large numbers of short-lived offspring: movement out of a location with high

population densities prevents overcrowding and decreases competition for resources. Dispersal is

somewhat localized, so that insects disperse from one resource to another within a limited

geographical range: for example, house flies and stable flies. Dispersal of insects may be similar

to diffusion of small particles in air, so that dispersal may be randomized, and may even go back

and forth between food/breeding resources during the insect's life-span (Schoof and Siverly

1954). In contrast, migration is typical of larger, longer-lived K-strategists that produce fewer

offspring. Migration generally involves longer distances, is uni-directional, and is a permanent

relocation. In insects, migration usually occurs over multiple life stages and/or generations: for

example, monarch butterflies. However, the differentiation between dispersal and migration can

be blurred, as is seen for locusts which migrate over long distances, with an environmentally

induced morphologically changed body, in response to limited food resources (Angelo and

Slansky 1984).

Although house flies are not thought to migrate, they do disperse frequently and readily.

Adult house fly behavior and dispersal is influenced by climatic factors (also referred to as

"environmental influences") such as temperature (Hewitt 1910), relative humidity (Hewitt 1910),

barometric pressure, light intensity (Greenberg 1959b), and electrostatic fields (Johnson 1969).

Dispersal of house flies and other synanthropic flies is of great interest, especially in livestock









and public-health situations, for both economic and medical reasons. Dispersal of house flies is

frequently studied through mark-release-recapture studies.

Mark-Release-Recapture Studies and Techniques

Mark-release-recapture (MRR) techniques are used to study insect dispersal, because

they provide point-to-point dispersal information about insect movement from a release site to a

recapture site. However, they do not describe the method or the path used (Turchin et al. 1991).

MRR studies have been conducted using different substances to mark the flies. Ideally, a marker

is not ubiquitous in the natural environment, but is quick and easy to apply, remains detectable

for a long period, does not adversely affect fly behavior or mortality, is inexpensive, readily

available, and has a long shelf-life (Turchin 1991).

Flies are marked using many different methods. In previous studies, flies have been made

radioactive (by digestion of P-32 radiolabelled diets) and been individually hand-painted, or

dusted with fluorescent dusts (Wong and Cleveland 1970). Another marking technique involves

amputating specific combinations of body parts and/or notching the exoskeleton in specific

locations. Of these techniques, fluorescent dusts are the most convenient to use (Hogsette, pers.

comm.), and best meet the above-listed requirements of an ideal insect marker.

Fluorescent dusts can be applied directly to adult flies, or the flies can self-mark (auto-

mark) themselves (Hogsette 1984). Auto-marking is accomplished by dusting a known breeding

site or by dusting flies in the pupal stage. Adult flies auto-mark themselves by direct contact with

the breeding substrate. Emerging flies will auto-mark themselves during emergence as the

ptilinum typically is dusted while emerging. However, detection of fluorescent dust on the

ptilinum is more difficult and time-consuming than detection of fluorescent dust on the

exoskeleton of flies that were marked as adults, because the ptilinum folds into the head capsule

after emergence. Therefore, detection of fluorescent dust on the ptilinum involves squeezing the









head capsule. Additionally, the quantity of dust on the ptilinum is typically much less than that

found on flies that are dusted as adults, further increasing the difficulty of using auto-marked

pupae.

Fluorescent dusts come in multiple colors which permits selection of colors that are best

for working with a particular insect. Day-GloTM arc-yellow and corona-magenta are the easiest to

use and detect with house flies (Hogsette, pers. comm.). Fluorescent dusts are sometimes visible

to the unaided eye, particularly if a great quantity is present on the insect, but these dusts are best

observed under long-wave ultraviolet (UV) light.

Flight and Dispersal

Flight is a dominating characteristic of most adult insects (Johnson 1969). Therefore, any

attempt to understand insect influence on disease transmission should incorporate learning about

the flight behavior and dispersal and/or migratory tendencies of the target insect. Insect dispersal

and migration is reviewed thoroughly by Johnson (1969). Insight into insect populations is

provided by Southwood (1966); his work has been revised so that newer editions are also

available. Methods to quantify insect movement are presented by Turchin (1998). Pedigo and

Buntin (1994) compiled the work of several authors to present a detailed resource for various

sampling methods applicable to agricultural, including livestock, insects.

Flight and dispersal behavior specific to house flies and related species of synanthropic

flies with similar breeding habits, such as stable flies and blow flies, has been reported by a

number of researchers and varies drastically in individual studies. In rural areas, house flies can

disperse 12 km (Broce 1993a) and have been documented dispersing up to 21 km (13 mi)

(Bishopp and Laake 1921, Alam and Zurek 2004) from their breeding sites. In urban

communities, most flies disperse within 1.7 km (1 mi) of release sites (West 1951, Quarterman et

al. 1954, Schoof and Siverly 1954bb; Hanec 1956; Sacca 1964, Milio et al. 1988). However,









house flies have been documented to disperse distances up to 33 km in urban environments

(Murvosh and Thaggard 1966). House fly dispersal speed has been documented at a rate of 1

km/h for the first 3-4 h, when dispersal occurred as direct flight over a large swampy area and

across rivers with 300-500 m widths in 24-50 h (Shura-Bura et al. 1962). Dispersal distances and

recapture rates might be influenced by the type of flies used, i.e., field-collected or laboratory-

reared. Previous studies have indicated that use of field-collected flies is more representative of

dispersal under natural conditions than flies that are reared for multiple-generations in the

laboratory. Eddy et al. (1962) recaptured a 10-fold higher percentage of field-collected flies than

laboratory-reared flies; this implies that laboratory colonies may lose the ability to disperse.

Dispersal of house flies increases the potential for transmission of zoonotic pathogens to

urban-residing humans, particularly from sites conducive to fly breeding such as dairies

(Kaufman et al. 2005, Ahmad et al. 2007, Conn et al. 2007), beef cattle feedlots (Skoda et al.

1993, Thomas 1993, Baldwin et al. 1996, Sanderson et al. 2006), swine facilities (Rosef and

Kapperud 1983, Halverson 2000), and poultry facilities (Hald et al. 2004, Watson et al. 2007). If

house flies can maintain a travel speed of 1 km/h for an extended period of time, and if house fly

dispersal flight occurs in a straight line from a breeding site, then house flies could potentially

transmit infectious pathogens as far as 12 km in only 12 h (Meerburg et al. 2007). The house fly

readily utilizes decomposing fecal/organic matter as well as human food, and moves freely

between the two. In locations where dairies and human communities are in close proximity,

house fly dispersal between the two could facilitate the transmission of enteric bacteria to

humans. External contamination of house flies can range from 2.5 to 29.5 million bacteria per fly

(Hawley et al. 1951), and some bacteria can survive up to 3.5 d on the surface of house flies

(Peppler 1944). Bacterial contamination of house flies can also occur after flies contact food









crops that have been fertilized with liquid slurry or solid fecal waste (Islam et al. 2005).

Mechanical transmission of bacteria by house flies has been well-established by many

researchers (Echeverria et al. 1983, Fotedar et al. 1992, Sasaki et al. 2000, Alam and Zurek 2004,

Buma et al. 2004, Ahmad et al. 2007, Nmorsi et al. 2007), whereas, biological transmission also

appears likely if E. coli 0157:H7 is capable of replicating within the house fly gut (Hawley et al.

1951, Petridis et al. 2006).

Fly dispersal has been measured by many types of mark, release and recapture studies

using fluorescent dusts, sticky traps, and UV lights (Hogsette 1983, Osek 2001). One of the

easiest and most efficient techniques for marking and releasing of large numbers of small insects

is the application of fluorescent dust (Hagler and Jackson 2001). Insects such as house flies are

collected in the field or mass-reared in the laboratory, marked for future identification, released

in the field, and recaptured at various distances from their release site.

Mark and Release Techniques

Many types of markers have been used effectively in the past, but are no longer

recommended or are sometimes prohibited under existing regulatory legislature, due to human,

animal and/or environmental health concerns. For example, radioactive phosphorus (32P) has

been added to adult fly laboratory diets; after successful feeding, 32P-labeled flies were released

and recaptured in the field (Lindquist et al. 1951, Yates and Lindquist 1952, Eddy 1962, Shura-

Bura et al. 1962). Marked flies were subsequently counted using readings on Geiger-counters

(Hoffman and Lindquist 1951, Lindquist et al. 1951, Yates and Lindquist 1952, Eddy 1962,

Shura-Bura et al. 1962). Hoffman and Lindquist (1951) reared flies in media containing 32P to

compare the efficacy of this method against application by ingestion, and determined that

feeding 32P to adult flies was both more effective and cost-effective. Lindquist et al. (1951)

compared marking adult house flies by adding 32P to the diet against dusting with fluorescent









dusts. They determined that marking with dietary 32P was more efficient and less labor-intensive,

and they observed that the dusts wore off within 48 h so that identification of dusted flies was

difficult. Similarly, Eddy et al. (1962) concluded that ingestion of 32P was a more useful marking

method than fluorescent dusts, because wild flies had natural fluorescence that was easily

confused with the fluorescent marker used in their study.

Although there is no universal method of marking insects, dusts are possibly the most

frequently used external markers, due to their ease of use in both application and observation, as

well as their low cost, ready availability, and low toxicity. Fluorescent dusts, in particular, Day-

Glo powdered pigment dusts, have been used to track dispersal and population dynamics,

without any observed adverse changes to insect behavior or mortality (Hogsette 1983, Hogsette

1984, Kristiansen and Skovmand 1985). Fluorescent dusts also offer potentially long-term

investigative study possibilities, because the dust has been shown to last up to 3.5 mo in the field

(Taft and Agee 1962). Flies are dusted with fluorescent powder, released, and recaptured;

subsequent UV light examination of recaptured flies illuminates any retained fluorescent dust on

areas of the body that the fly has difficulty grooming. Flies dusted as adults will typically retain

dust particles on portions of the thorax; when the puparia are dusted, flies emerge, crawl through

the dust, and retain it on their ptinilum (Hogsette 1983).

An additional advantage of fluorescent dusts is that their visibility is greatly enhanced

when examined under long-ultraviolet (UV) light. Thus, large numbers of recaptured house flies

on sticky traps can be examined rapidly and easily under UV light to determine how many are

marked. This eliminates time- and labor-intensive observation methods used with alternative

marking techniques, as there is no need to destroy individual insects to observe internally

expressed dyes, to apply solvents, or to perform genetic analysis. Application of fluorescent









dusts is relatively easy, inexpensive, and less labor-intensive than other insect marking

techniques, and enables marking of thousands of insects simultaneously (Zhao et al. 1999).

Additionally, application of fluorescent dusts can be accomplished using mechanical dusters

(Hogsette et al. 1993).

Population Dynamics: Monitoring House Fly Populations

Fly populations at dairy barns typically need to be monitored in order to determine the

effectiveness of control methods or to measure seasonal and weather-related fly population

changes. There are many different methods available to monitor fly populations; each has its

strengths and limitations. Ideally, a fly monitoring method will provide sensitive, accurate

results, be easy to use and interpret even by inexperienced persons, be inexpensive, require very

little investment of time and labor (Pickens et al. 1972), be protected from cattle (Morgan and

Pickens 1978), and not interfere with cattle operations or endanger cattle health and safety.

Monitoring devices must be placed out of reach of cattle, particularly if they are small enough to

be eaten by cattle. This is particularly true of spot cards, described further below. Fly monitoring

can be conducted either by passive or active methods.

Passive monitoring methods

Passive methods of monitoring dairy barn fly populations are those which use one or

more of the following: spot cards, sticky traps, ultraviolet (UV) light traps, and baited traps

(Morgan and Pickens 1978). Spot cards are plain white cards (8 x 13 cm) that are conveniently

sized for easy transport and use (Axtell 1970, Lysyk and Axtell 1986), but they alternatively may

be of larger dimensions such as 13 x 20 cm cards (Pickens et al. 1972). Standard office-supply

index cards are often used as spot cards. Regardless of size, spot cards are placed vertically or

horizontally flush against barn walls, beams, or rafters (Lysyk and Axtell 1985), in areas where

house flies rest. Spot cards offer many advantages for monitoring of fly populations. They are









convenient, economical, easy to transport, readily available, and can be used by relatively

untrained personnel, including busy farmers.

Flies continuously regurgitate and excrete while resting, so spot cards provide a

convenient method to obtain house fly relative abundance data in dairy barns when placed in

house fly resting areas. Spot cards provide useful information about the changes in house fly

populations from week to week, and help ensure optimal fly control efforts, because the success

of fly treatments can readily be ascertained. If the number of house fly spots decreases, then the

treatment decreased the fly population. Successful treatment would be defined by reducing the

number of spots per card below some pre-defined threshold that often is farm-specific.

Spot cards may also be suspended or hung vertically from barn rafters, however, this does

not appear to provide useful results as Pickens et al. (1972) did not find spot cards sensitive to

changes in house fly population density. They observed that the number of fecal and

regurgitation spots remained relatively the same despite an artificially doubled house fly

population due to releases of marked flies within an enclosed barn. However, they hung their

spot cards vertically from the ceiling rather than placing them flush against the wall as reported

by Lysyk and Axtell (1985). House flies prefer to rest along straight edges. Therefore, hanging

large cards from the ceiling was probably not ideal, especially since Pickens et al. (1972)

described the presence of a large exhaust fan at one end of the barn. Presumably, the fans created

air currents that might have inhibited fly resting on their spot cards, particularly if their spot

cards were moving in an air current. Furthermore, Pickens et al. (1972) released laboratory-

reared flies, and there is some evidence that laboratory-reared flies do not take flight as readily as

flies from natural populations (Eddy et al. 1962). Thus, the flies used in their experiment might

not have dispersed throughout the enclosed dairy barn as readily as native flies.









In addition to monitoring house fly population changes when placed correctly, spot cards

also might permit isolation of fecal and/or regurgitation spots for microbial or genetic analysis of

digestive tract contents, although they do not appear to have been used for this purpose yet. In

dairy barns that have multiple species of flies, fecal and regurgitation spots could be due to any

of the various species present. However, judicious placement can reduce species overlap by

placing cards where house flies are observed resting. Because house flies are not actually

captured on spot cards, individual fly species cannot be identified from spot cards using standard

methods. However, if fecal and vomit spots contain any host cells from the flies, then the

potential exists for identification of particular fly species by genetic analysis such as polymerase

chain reaction (PCR). Feces are typically surrounded by a layer of peritrophic membrane and

excreted as discrete "fecal pellets." Therefore, host cells should be present with the fecal spots,

so that the potential for identification of fly species by analysis of spot cards offers an interesting

and promising possibility.

Sticky cards (Hogsette et al. 1993) provide another useful house fly population

monitoring method. Like spot cards, they are easy to transport, easy to use by untrained

personnel, and inexpensive. Fresh cards are recommended to achieve optimal results, because the

adhesive coating can dry out over time. An advantage that sticky cards offer is the ability to

easily identify captured flies to species. Flies become stuck on the adhesive and are therefore

readily available for examination, either in the field or in the laboratory. Like spot cards, sticky

cards offer the potential for genetic analysis. Because the entire fly body is present, genetic

analysis could be used to determine the genetic makeup of the flies (for phylogenetic studies) or

to analyze the gut contents. Although sticky cards do not appear to have been used for either

purpose, they potentially provide another tool for increased understanding of house fly









population dynamics as well as determination of microorganism survival on and within the house

fly. Preuss (1951) obtained viable Salmonella sp. from a house fly seven days after it was killed,

so the potential for isolating bacteria from house flies captured on sticky cards is strong. A

disadvantage of sticky cards is that they can become ineffective if conditions are dusty, because

the adhesive becomes covered with dust and debris, so that flies do not adhere after landing on

the card. Attempts to correlate spot card and sticky counts have been variable. Turner and

Ruszler found no correlation, while Geden et al. (1999) found a high correlation.

Originally used to capture and monitor stable flies, alsynite traps (Williams 1973) have

proven effective for capturing and monitoring house flies and other species of insects. The

fiberglass panels reflect sunlight in plane-polarized ultraviolet wave lengths, making them

attractive to many flying insects: e.g., stable flies (Williams 1973), alate red imported fire ants,

Solenopsis invicta Buren, (Milio et al. 1988), and house flies (Geden 2006). The original alsynite

trap consisted of translucent rectangular fiberglass panels coated with an adhesive (Williams

1973). Berry et al. (1981) modified the trap by placing adhesive-coated sleeves on the

interlocking panels instead of applying adhesive directly to the alsynite panels. The sleeves can

be removed and taken to a laboratory where insects can be examined and counted. Alsynite traps

have proven to be effective, easy to transport, and easy to use in the field. A modified alsynite

trap (Broce 1988) consists of a translucent rectangular fiberglass panel wrapped to form a

cylinder with an adhesive coated clear plastic sleeve that wraps around the outside of the

cylinder for easy removal and replacement in the field.

Active monitoring methods

Active methods of monitoring house fly populations include visual counts, Scudder grids

(Scudder 1947, 1949; Pickens et al. 1972), and sweep netting (Morgan and Pickens 1978). Non-

biting fly population densities can be estimated for some species by active visual counts. For









example, Fannia canicularis (L.), the lesser house fly, has a behavioral habit of hovering in

sunny areas. Therefore, one can estimate a lesser house fly population by estimating the number

of flies hovering in a sunlit circular field 2 m in diameter, by counting in increments of 10

(Morgan and Pickens 1978). Visual observations invariably are estimations, and can be biased

due to researcher experience and ability to recognize house flies versus similar-appearing

muscids. A disadvantage of visual observation is that flies cannot be identified, sexed or aged, so

that the population dynamics remain unknown.

Scudder grids (Scudder 1947, 1949) are another active monitoring method. They provide

instantaneous "snapshot" pictures of house fly population densities, and are very useful for

monitoring changes in activity levels and population densities. Scudder grids are particularly

effective for monitoring the success of control efforts if used repeatedly at the same physical

location both before and after treatment (Scudder 1947). Placement of the grids is critical.

Because house flies are not distributed randomly throughout a dairy, but aggregate at food and

breeding sites, Scudder grids should be placed in areas of highest fly density. Scudder (1947)

found that results were most consistent with observed population trends and most consistent

from week to week when he obtained multiple counts at each site and recorded the average of the

highest three counts. Some disadvantages of the Scudder grid are that researcher proximity,

especially if casting a shadow over the grid ; or repeated placement of the grid in one spot within

a short period of time, which can decrease Scudder grid counts, because the flies move away

from the disturbance. Due to fly responses to this monitoring method, Scudder grid counts do not

necessarily correlate to an observable population of house flies for the purposes of quantifying

population densities. Grids should be placed by the same person to decrease user-induced

variability influences on fly counts. As with visually observed flies, flies that land on Scudder









grids cannot be identified, aged or sexed, so information regarding the population dynamics

remains unknown. However, for the experienced person, Scudder grids can provide an

inexpensive, easy-to-use and reliably repeatable method for monitoring increases or decreases in

fly populations. Their ease-of-use and portability in the field make them useful monitoring

methods at dairies and other animal-rearing facilities.

Sweep netting at dairies is another fast and easy method used to monitor adult house fly

populations. Like Scudder grids, sweep netting is performed on-site to gain an instantaneous

"snapshot" picture of house fly and other flying insect population densities (Dhillon and Challet

1985, New 1998). However, unlike the Scudder grids, repeated sweep netting reduces the

available population, because one is removing individuals from the environment and greatly

disrupts flies in a given area. In contrast, Scudder grids do not directly impact population sizes

and minimally impact flies in a given area. Sweep netting would not noticeably impact high-

density populations; however, when fly populations are low, sweep net counts for successive

sweeps may decrease with each sweep. Like the Scudder grid counts, sweep nets should be

performed in fly aggregation areas, at the same time of day, and by the same person. Sweep net

counts can vary dramatically between collecting individuals, based on experience and technique.

It might be important to note that sweep netting captures only airborne flies, and flies are

typically captured by purposely disturbing adult flies that are resting, feeding or ovipositing.

Therefore, sweep netting might not provide accurate information regarding the normal level of

airborne activity. However, an advantage of sweep netting over other monitoring methods is the

capture of live adults, so that analysis of population ratios can be made. Collected flies can be

identified to species, sexed and aged and female flies can be examined for reproductive status.

Collection of mostly young adults 1-3 d old would provide information that the flies had only









recently emerged. Because sweep netting enables the researcher to obtain information regarding

population dynamics, it is a useful monitoring method at dairies and other animal-rearing

facilities.

House Fly Economical Impacts and Disease Outbreaks

House flies are strongly associated with humans and livestock. This synanthropy, when

combined with the house fly's caprophagic and saprophagic eating habits, makes the house fly an

important vector of human diarrhegenic pathogens. Diseases which are transmitted via the fecal-

oral route are especially prone to dissemination by the house fly. In locations where food is being

prepared for consumption, such as restaurants, the potential for bacterial contamination by the fly

is tremendous, as the fly travels freely between decomposing organic matter found in restaurant

garbage dumpsters, exposed kitchen surfaces and foods, dining tables, and even restaurant

bathrooms. The bacterial diversity and quantity on restaurant-associated house flies has been

examined (Nayduch et al. 2001, Butler et al. 2010), especially at ready-to-eat (RTE) food

establishments (Macovei and Zurek 2008).

Diarrheal diseases impart an enormous economic toll on human and agricultural animal

(e.g., cattle) populations, with severe health and economic impacts. Hospital expenses for human

patients with infectious diarrhea can four times greater than for other patients. Similarly,

medication expenses can be four times higher and the length of hospitalization can be three times

longer (Suda et al. 2003). Economic impacts include lost income for families that must miss

work, as well as lost profits for employers (CDC 2002). For cattle, increased operating expenses

are incurred by dairy farms, feedlots and cattle rendering plants that must increase fly and

pathogen surveillance and management measures to comply with federal regulatory mandates

(CDC 2009). Indirect losses include increased labor costs associated with fly control and

sanitation efforts, which may include increased fuel costs to operate composting and waste-









removal equipment, increased water costs to wash barns and clean equipment, increased

expenses for insecticides and traps, and additional wages to employees who perform all the

additional tasks (Lazarus et al. 1989). It is interesting to note here that expenses related to

insecticides can increase quickly because of the house fly's rapid development of insecticide

resistance. House fly insecticide resistance can occur quickly enough to necessitate increased

usage of insecticides, thus driving up the expense dramatically. Alternatively, one may switch to

a pesticide containing a different active-ingredient; however, this still results in overall increased

expenditure towards the effort of controlling fly populations in an effort to prevent diarrheal

disease occurrence.

Additional indirect losses can include legal expenses and forced farm closures (Thomas

and Skoda 1993). Courts can impose stiff fines upon many different food-related industries

including producers, distributors, and restaurants. A recent example: in June, 2008, a lawsuit led

to a $13.5 million settlement after a child in Milwaukee, Wisconsin died due to consumption of

E. coli 0157:H7-contaminated food (Powell, 2008).

House Fly Management

Pest control efforts include cultural, mechanical, biological, and chemical methods. Each

method is discussed below, and the integration of these methods into one comprehensive control

program known as Integrated Pest Management (IPM).

Cultural control

Cultural control includes sanitation and management efforts, e.g., moisture and manure

removal at dairies designed to prevent fly populations from building to unacceptable levels

(Stafford 2008) by eliminating immature fly development areas. Flies breed in spilled feed, moist

hay and manure. Weekly removal of these materials is recommended (Rutz et al. 1994). Spilled

feed and manure can accumulate and retain moisture under fence edges, along the sides of liquid









manure pits, and in other hard-to reach areas (Rutz et al. 1994, Farkas and Hogsette 2000).

Preventing moisture in fly breeding areas, e.g., in spilled grains, is an important part of cultural

control (Farkas and Hogsette 2000). Watering devices should be maintained to prevent leaks, and

manure should be removed to maximize drainage. Flies can breed in manure that lines manure

pits or floats on the surface of manure pits and lagoons. Therefore, cultural control of fly

breeding sites includes preventing manure clumps from lining manure pits or from floating on

the surface (Farkas and Hogsette 2000). Without adequate removal of manure and other fly

breeding sources, chemical efforts to control the house fly population will be less effective and

more expensive (Farkas and Hogsette 2000). Cultural control methods also encourage the growth

and development of house fly predators and parasites (biological control).

Additional cultural control methods include using screens as exclusion barriers on

windows and doors, preventing access to garbage, and composting garbage, manure and soiled

bedding properly so that decomposition is aerobic and hot enough to kill developing flies

(Stafford 2008).

Mechanical control

Mechanical control methods use non-chemical devices to kill flies that are in the

environment. Fly swatters provide low-tech mechanical control of individual house flies

(Stafford 2008). Low numbers of flies can be controlled by using sticky traps, jug or cylinder

traps and bag traps. Sticky traps are coated with adhesive materials, so they are less effective

when fly numbers are high or when environmental conditions are dusty (Stafford 2008). Sticky

traps are most effective when used indoors; however, they cannot be used in food-preparation

settings because become unsightly and they can drip when temperatures become too warm

(Carlson and Hogsette 2007). Fluorescent or ultraviolet light emitting electrocution traps are

useful for mechanically controlling larger fly populations (Stafford 2008). The flies are









electrocuted when they contact the electric mesh. However, electrocution of insects results in

airborne-scattering of insect parts and infectious microorganisms (Pickens 1989, Ananth et al.

1992, Broce 1993b, Tesch and Goodman 1995, Broce and Urban 1998). Therefore, use of

electrocuting traps is not recommended, and is sometimes prohibited, in medical and food-

preparation settings. An additional disadvantage of electrocuting grids is that they kill high

numbers of non-target insects (Frick and Tallamy 1996). Many mechanical traps use ultraviolet

light traps with sticky glue boards instead of electrocuting grids, although the attractiveness of

these traps relative to the attraction of food in the food-preparation area is unknown (Carlson and

Hogsette 2007). Bulbs in light traps lose their effectiveness over time, so that they become less

attractive to flies. Thus, it is important to replace bulbs on a timely basis. Additionally, indoor

traps should be installed so that they attract flies that are already in the local environment, rather

than attracting flies from far away into the establishment.

Some mechanical control traps use attractant baits that draw house flies into a non-toxic

solution (Stafford 2008). Examples include jug traps, bag traps, and metal or plastic cylinder

traps. Baited traps are useful for controlling large numbers of house flies; some use only water as

the bait, while others add proprietary chemical mixtures that mimic natural food materials or

pheromones, such as the female house fly sex attractant (Z)-9-tricosene. Because (Z)-9-tricosene

is odorless to humans, it is a useful indoor bait. In contrast, baits that mimic natural food sources

typically have strong unpleasant odors and are therefore best suited for outdoor use. Bait traps

come in a wide variety of styles and sizes, with some designed to be hung and others designed to

be placed on the ground. In general, bait traps take advantage of the house fly's positive

phototrophic behavior by providing small access holes through which the flies can enter the trap,

but not exit. Therefore, the flies die inside the trap. Jug and cylinder (container) bait traps can be









reused by emptying them when approximately one-third full, and rebaiting, so that they provide

an economical control method. Homemade bait traps are easily made from milk cartons and

plastic soda bottles by cutting the top portions off and replacing them in an inverted position to

acts as a funnel. Flies will readily enter the carton or bottle through the funnel, but will not exit.

Like bag traps, carton and bottle traps are disposable (Stafford 2008).

Biological control

Biological control of house flies is achieved through predation or parasitism by natural

house fly enemies or by infection with pathogens that kill the house flies. While biological

control occurs unassisted in the natural environment, attempts to amplify its actions through

release of increased biological control agents are ongoing. Natural enemies of house flies include

other arthropods, fungi, bacteria, and viruses (Barnard 2003, Geden 2006, Stafford 2008).

Parasitoid wasps used for biological control of house flies are often species in the family

Pteromalidae, from the following genera: Spalangia, Muscidifurax, and Nasonia (Morgan et al.

1979, Crespo et al. 1998, Tobin and Pitts 1999, Floate et al. 2000, Kaufman et al. 2001a, Geden

2006, McKay et al. 2007, Birkemoe et al. 2008). One benefit of the parasitoid wasps is their

relative target-specificity, due to the postulated co-evolution of the parasitoids with their

synanthropic muscoid hosts (Pimentel et al. 1963). Some species of wasps will target both house

flies and stable flies, which are two of the most economically-damaging fly species in livestock

environments. Efforts to identify new species of parasitoid wasps, and to commercially rear them

are ongoing.

Six hymenopteran parasitoids that specifically attack fly pupae were evaluated by Geden

(2006): Muscidifurax raptor Girualt and Sanders, Spalangia cameroni Perkins, Spalangia

nigroaenea Curtis, Spalangia endius (Walker), Spalangia gemina Boucek (Hymenoptera:

Pteromalidae), and Dirhinus himalayanus (Hymenoptera: Chalcididae). Hogsette et al. (2001)









obtained seven hymenopteran pupal parasitoids from house fly pupae in Hungary: Spalangia

cameroni Perkins, S. nigroaenea Curtis, S. endius Walker, Pachycrepoideus vindemiae Rondani,

Trichomalopsis sp., and two undetermined spp. of Diapriidae. Other types of wasps reported as

parasitoids of house flies include species in the family Ichneumonidae: for example, Exeristes

comstockii (Cress) (Hymenoptera: Ichneumonidae) (Bracken 1965). Hogsette et al. (2001)

reported an undescribed species of Brachymeria parasitizing a house fly pupa in Hungary.

Some dipteran species are also useful for biological control of house flies. Some are

predators, such as the bronze dump fly (Byrd and Castner 2009), sometimes called the black

garbage fly, Hydrotaea (=Ophyra) leucostoma (Weidemann) (Diptera: Muscidae) (Anderson and

Poorbaugh 1964), and the black dump fly, Hydrotaea (= Ophyra) aenescens (Weidemann)

(Diptera: Muscidae) (Hogsette and Jacobs 1999, Hogsette et al. 2002). Hydrotaea larvae actively

pursue and attack house fly larvae, but rarely the pupae (Anderson and Poorbaugh 1964).

Hydrotaea eat the visceral tissues, but not the larval integument or puparium. One Hydrotaea fly

can kill up to 20 house fly larvae in one day (Anderson and Poorbaugh 1964). Hogsette and

Washington (1995) developed a method to mass-rear Hydrotaea aenescens for biological control

studies.

Other Diptera are effective biological control agents by non-predatory methods. One such

example is the black soldier fly, Hermetia illucens (L.) (Diptera: Stratiomyiidae) (Furman et al.

1959; Sheppard 1983; Sheppard et al. 1994; Sheppard et al. 2002). Studies indicate that soldier

fly larvae outcompete house fly larvae for food resources in the breeding habitat, and, in this

manner, limit house fly growth and larval development (Sheppard 1983).

Bacteria such as Bacillus i1/ui ilgienl\i\ (Zhong et al. 2000, Ruiu et al. 2007) and

Brevibacillus laterosporus (Ruiu et al. 2006) have been reported as useful for biological control









of house flies, caterpillars and mosquito larvae (Swadener 1994), due to production of

insecticidal endotoxins (Zhong et al. 2000; Ruiu et al. 2006, 2007). Bacillus thuringiensis subsp.

israelis (Bti) produces a delta-endotoxin (Zhong et al. 2000, Ruiu et al. 2006). However, there is

evidence that Bti and B. laterosporus are toxic against the house fly parasitoid, M. raptor,

although much less so than against the house fly (Ruiu et al. 2007).

Fungi are considered by some researchers to show promise as biological control agents

against house flies (Mullens and Rodriguez 1986, Geden et al. 1993, Steinkraus et al. 1993,

Watson and Petersen 1993). Two fungi that successfully infect and kill adult house flies are

Entomophthora muscae (Chon) Fresenius and Beauveria bassiana (Balsamo) Vuillemin.

However, these fungi are limited to specific climatic conditions, and are dependent upon high fly

population densities (Watson and Petersen 1993). Although these fungi difficult to produce

commercially well enough to be effective and economically viable commercial house fly

biological agents (Geden et al. 1993, Watson and Petersen 1993), recent rearing techniques have

resulted in a commercial product that is highly effective (Kaufman et al. 2005a). Beauveria

bassiana has proven to be effective against both adult and larval house flies (Watson et al. 1995).

Chemical control

In general, chemical control of house flies has been the most widely used approach over

the past 60 years. Insecticides are applied in a variety of ways, including space sprays, residual

wall sprays, feed through products, on-animal applications, misting systems and toxic fly baits

(Rutz et al. 1994). Pyrethrin fogs and space sprays are recommended for initial use because they

work well in conjunction with biological control for integrated pest management (Kaufman

2002). Insecticides can adversely affect biological control agents, particularly if used early in the

season because parasite populations lag behind house fly populations (Rutz et al. 1994).

Elimination of biological control agents can result in an increased dependence on insecticides









that can increase the development of insecticide resistance in the house fly population (Rutz et al.

1994). Residual pesticides are recommended only for use emergencies and for use late in the

season (Kaufman 2002). Toxic fly baits cannot be used in food establishments, including milk

storage rooms on dairies, or in areas where children are present. They also should be avoided in

areas frequented by children or pets, and must be placed in a container but not sprinkled on the

ground.

Resistance to chlorinated hydrocarbons develops more quickly in larvae than in adults

(Miles 1959). He concludes that chemical control should therefore be restricted to adults, to

minimize development of resistant fly strains. However, adult flies can develop tremendous

resistance against over-used pesticides: Cao et al. (2006) reported 13- to 250-fold greater

deltamethrin resistance in adult house flies collected from urban garbage dumps in Northern

China than from laboratory susceptible strains.

Integrated Pest Management

Integrated pest management incorporates a combination of several available fly control

methods (Kaufman et al. 2002). Each control method that is used to control fly populations at

animal-rearing facilities such as dairies is insufficient or uneconomical if used alone. The

foundation of any successful IPM program is sanitation and manure management. Without

adequate cultural controls, neither biological nor chemical controls will be effective.

Use of biological control decreases dependence upon chemical insecticides (Farkas and

Hogsette 2000). However, natural parasitoids used for fly control differ in efficacy depending

upon geographical region, season, climate, habitat, host density and host distribution. Thus, it is

important to use naturally-occurring species of parasitoids for a particular area, and it is vital to

understand both the house fly and parasitoid bionomics to effect a successful long-term

biological component of the overall IPM program (Farkas and Hogsette 2000). Parasitoid









populations can be reduced below effective levels by indiscriminate use of broad-spectrum

insecticides used to control house fly populations, and by manure removal. Mass releases of

parasitoid wasps are often necessary to reduce house fly populations throughout the season,

because the parasitoids need to be consistently present in the manure. For this reason, although

sanitation is the first step in successful fly control, manure removal should not be complete, but

should leave a base of old manure for house fly parasites and predators (Farkas and Hogsette

2000).

Chemical control can achieve a rapid reduction in adult flies, but is limited in use due to

side effects or poisoning when overused, to breed sensitivity, to interactions with medication

administered to livestock animals, to contamination of food products such as milk, and to the

development of resistance among the fly populations (Farkas and Hogsette 2000, Pimentel

2002). Therefore, chemical control should be used in conjunction with cultural/sanitation,

mechanical, and biological control methods (Rutz et al. 1994, Farkas and Hogsette 2000,

Kaufman et al. 2002).

Usefulness of House Flies

House flies are useful in some ways that might be unexpected. For example, house fly

pupae have been used to achieve relatively species-specific chemical control of several ant

species, including Solenopsis invicta Buren, the red imported fire ant (RIFA). Ants are extremely

effective predators of house fly larvae. Williams et al. (1990) administered hydramethylnon- and

fenoxycarb-coated house fly pupae to eight species of ants. They achieved 100% mortality in

three species (including RIFA) and <20 % mortality in the remaining five species by using house

fly pupae as bait carriers. They note that use of house fly pupae permits more selective control

over species that consume house fly pupae, and that mass-production of the house fly pupae

could make this potential method of ant control more economical (Williams et al. 1990). For









baits, house fly pupae was not practical. However, house fly pupae treated with cyclodienes were

used for aerial applications in parts of the United States.

House flies are potentially useful in an interesting agricultural application to help prevent

browning of apples. House fly pupae contain high levels of an apple polyphenol oxidase

inhibitor that inhibits adverse browning in apple and apple products (Yoruk et al. 2003).

The house fly might be a promising livestock and poultry food resource (Eby and Denby

1978, Boushy 1991) due to the very aspects of the house fly's biology and behavior that make it

a global pest. The house fly converts decomposing organic matter and fecal materials into body

tissue during its larval stages. In doing so, it reduces waste, generates energy and creates a high-

nitrogen food resource. House flies have been used as feed supplements for swine (Poluektova et

al. 1980, Chiou and Chen 1982) and poultry (Calvert et al. 1969, Papp 1974, Teotia and Miller

1974, Gawaad and Brune 1979). Freeze-dried fly larvae are also commercially available for use

as fish feed.

However, the use of immature flies as feed supplements might not be advisable unless

they can be subjected to bactericidal conditions, because there is evidence that animals can

become infected with pathogenic bacteria present in the consumed flies (Gerberich 1952). Even

if flies are not used as a feed resource, they are still useful because they perform the beneficial

act of decomposing manure (Miller et al. 1974, Beard and Sands 1973). This could reduce the

amount of waste and decrease labor costs associated with waste-management efforts.

Although not intentionally provided as supplemental feed, adult flies might be consumed

by animals, particularly poultry, after they are killed by pesticide treatment at an animal-rearing

facility. Therefore, efforts to reduce consumption of flies killed by pesticides might be advisable.

Greenberg (1959e) postulates that consumption of adult flies might result in pathogen









transmission after citing the work of Gross and Preuss (1951) in which they determined that

viable Salmonella sp. were recovered in flies 7 d after they were killed with DDT. Survival of

pathogenic bacteria in dead insects illustrates the importance of sanitation in conjunction with fly

control in order to reduce disease transmission potential at animal-rearing facilities.

Enterobacteriaceae

Enterobacteriaceae Rahn 1937 is a family of opportunistic, facultative anaerobic, non-

spore-forming, gram-negative, rod-shaped bacteria, found as normal commensal fauna in healthy

human and domestic animal gastrointestinal tracts (Caprioli et al. 2005, Madigan and Martinko

2006, FDA-CFSAN 2007a). Enteric microbiological communities tend to be very complex, with

bacterial populations of more than 1014 cells, which is ten times more cells than the number of

cells that constitute the human body (FDA-CFSAN 2007a). Enteric bacteria belong to more than

500 different species (Steinhoff 2005), which collectively perform various metabolic activities,

such as digestive fermentation and vitamin synthesis upon host-consumed nutrients (FDA-

CFSAN 2007a). With this level of complexity, resultant fecal bacteria concentrations can exceed

1012 cells/g of feces (Couteau et al. 2001).

Some enterobacteria are opportunistic pathogens that cause diarrhea, dysentery,

meningitis, typhoid fever, and food poisoning (MEDIC 1995) to mammalian hosts through a

variety of mechanisms (Manafi 2003). Disease-causing enterobacteria include several closely

related genera: Escherichia, iigell/, Salmonella, Yersinia, Klebsiella, Proteus, Edwardsiella,

Citrobacter, Enterobacter, Serratia, Providencia, and Morganella (FDA-CFSAN 2007a).

Although Escherichia is listed at the genus level with the above group, it is unique due to the fact

that only one of its seven species (Euzeby 2008, NCBI. 2008), E. coli, is known to be pathogenic

to mammals. In contrast, other genera have several pathogenic species.









In addition to shared morphological and physiological traits, members of the

Enterobacteriaceae share biochemical characteristics that make differentiation and identification

difficult: all ferment glucose, reduce nitrates, and are oxidase-negative. Despite these

similarities, other biochemical characteristics are diagnostic for differentiation. For example,

lactose fermentation is typical of non-pathogenic enterobacteria, while non-fermentation of

lactose is typical of pathogenic enteric bacteria. Selective and/or differential media used in the

laboratory to identify pathogenic strains takes advantage of this lactose-fermentation

characteristic by inclusion of lactose and dyes which provide presumptive identification by easily

observed color changes where fermenting and non-fermenting bacteria will produce colonies of

distinctly different colors.

Among the Enterobacteriaceae, the genus Escherichia is comprised of seven species

(Euzeby 2008, NCBI 2008), one of which is E. coli. Pathogenic strains of E. coli present health

risks, partially because of their low infectious dose, which is variously reported as less than 100

organisms/g of feces (USDA-FSIS 2008a, USDA-FSIS 2008b), less than 50 organisms/g of feces

(Tilden et al. 1996), and possibly as few as 10 organisms/g of feces (FDA-CFSAN 2007b).

Escherichia coli

Escherichia coli was originally described in 1885 as Bacterium coli commune, by

Theodor Escherich (Janda and Abbott 2006, NCBI 2008, Todar 2008). An additional synonym of

E. coli used in older literature is Bacillus coli Migula 1895 (NCBI. 2008). Esherichia coli is the

most abundant, commensal, facultative anaerobic bacteria in many mammals, including humans.

(Donnenberg 2002). Homeothermic animals are the natural reservoir of E. coli, and E. coli is part

of the normal gut fauna in nearly all domestic animals (Bettelheim 1991), and accounts for

approximately one percent of fecal biomass (Janda and Abbott 2006). Human stool contains a

diverse bacteria fauna of 1011 cells per cubic centimeter, with 109 E. coli per cubic centimeter









(Neidhardt et al. 1996). Escherichia coli is a rod-shaped bacterium with an approximate length of

2.5 rm and diameter of 0.8 rm (Berg 2004). In contrast, a human hair is 80 pl in diameter

(Neidhardt et al. 1996). When placed in warm nutrient broths, E. coli replicates in 20 min (Berg

2004).

Commensal E. coli appears to produce cofactors and to inhibit pathogenic colonization in

the digestive tract (Donnenberg 2002). Escherichia coli is comprised of more than 700 strains

(Todar 2008) and more than 200 of these strains produce Shiga toxins (Thorpe et al. 2002), of

which most of those strains are pathogenic (Madigan and Martinko 2006, FDA-CFSAN 2007a).

All pathogenic strains are enteric (Madigan and Martinko 2006), but the pathogenic potential of

a particular strain depends on the specific collection of virulence genes (Donnenberg 2002).

Within the anaerobic gastrointestinal tract of animals, non-pathogenic commensal strains

of E. coli synthesize vitamins and inhibit growth of pathogenic microorganisms (FDA-CFSAN

2007c). Both behaviors benefit the mammalian host with improved nutritional uptake. In

humans, E. coli is the predominant bacterium in the large intestines (FDA-CFSAN-2007a),

whereas in cattle, E. coli is a normal component of the rumen, the largest chamber of the four-

chambered stomach (Callaway et al. 2004). Although consistently present in all animal

gastrointestinal tracts, E. coli prevalence, density, survival and fecal shedding rates can differ

among hosts. For example, cattle shed E. coli 0157:H7 intermittently (Zhao et al. 1995, Shere et

al. 1998). Differences in host diet have been implicated in increased fecal shedding of E. coli

0157:H7. For example, E. coli from grain-fed cattle are especially resistant to low pH and have

greater survival rates in acidic enteric environments than E. coli from forage-fed cattle (Callaway

et al. 2003).









Outside the host's facultative anaerobic intestinal environment, E. coli can be

outcompeted by other species of bacteria. Host excretion of manure rapidly exposes E. coli to

multiple systemic shocks. The two most important shocks relate to transition: 1) from an ideal

370 C to ambient air temperature, and 2) from an anoxic environment to a highly oxygenated

environment, which is relatively toxic to E. coli (Doyle and Beuchat 2007). Separately, either of

these two shocks can result in serious, but reversible, injury to E. coli; however, when combined,

they can cause irreversible damage and mortality.

Because E. coli is a facultative anaerobe, the aerobic environment is not directly lethal to

E. coli. Instead, the aerobic environment presents an indirect, potentially more harmful, obstacle

to E. coli's survival outside of the enteric environment: competition with aerobic

microorganisms. Competing organisms can inhibit E. coli directly by secreting antibiotics or

toxins (Xavier and Russell 2006), or indirectly by consuming available nutrients (Hibbing et al.

2010).

Concentrations of E. coli in hosts differ among host animal species. Within a particular

animal, concentrations also can differ depending on gastrointestinal location. In cattle,

concentrations are high in the mucosal-anal region because it is the site ofE. coli 0157:H7

attachment (Greenquist et al. 2005). In cattle, prevalence and density ofE. coli in fecal samples

varies widely, and may be influenced by factors including diversity of the bacteria community

within the gastrointestinal tract, diet, season, lactating stage (Hancock et al. 1994), and

application of vaccines (Van Donkersgoed et al 2005). Many different strains of E. coli may be

present in one individual simultaneously, as seen in humans (Beutin et al. 2004) and cattle

(Majalija et al. 2008).









There is some level of host specificity between certain strains ofE. coli and a particular

animal host. While commensal bacteria serve a useful, and essential role in digestion for the host

animal, these bacteria can become pathogenic if they are introduced into a different animal

species. For example, E. coli 0157:H7 is a commensal bacterium in both livestock and wild

ruminants. However, this strain ofE. coli is a human pathogen that can result in severe illness

and death. In humans, E. coli 0157:H7 causes diarrhea, dysentery, hemolytic colitis (HC), and if

left untreated, can lead to development of hemolytic uremic syndrome (HUS) and acute kidney

failure (Tarr and Neill 2001).

Pathogenic Escherichia coli

Although most E. coli are beneficial to their associated host, a subset ofE. coli are

pathogenic, and can cause diarrheal diseases (FDA-CFSAN 2007b). Escherichia coli is a clonal

species, with clones differentiated into serotypes based on various combinations of somatic (0)

and flagellar (H) antigens (Wang and Reeves 1998). The O and H antigens are most frequently

used for serotype identification and differentiation. There are at least 181 E. coli 0 antigens

(Durso et al. 2007). Many of these antigens are associated with pathogenicity (Wang and Reeves

1998) whereby these O antigens are considered virulence factors (Wang and Reeves 1998).

There are approximately 200 pathogenic strains ofE. coli, and they are broadly grouped into

large subsets: pathogenic E. coli (FDA-CFSAN 2007b), enterovirulent E. coli FDA-CFSAN

2007b), diarrheagenic E. coli (Nataro and Kaper 1998, FDA-CFSAN 2007c), and

enterohemorraghic E. coli (FDA-CFSAN 2007b). There is overlap between some of the groups,

because E. coli strains are placed into groups based on possession of particular virulence factors

such as O and H antigens and biochemical traits. Therefore, serotypes that possess multiple

virulence factors sometimes fall into more than one group. In general, diarrheagenic E. coli are a

subset of pathogenic E. coli, and enterohemorrhagic E. coli are a subset of diarrheagenic E. coli.









Pathogenic E. coli include all of the disease-causing strains, regardless of etiology. Some

of the more infectious and therefore most studied O antigen serogroups include: 026, 055, 086,

0103, O11 lab, 0119, 0125ac, 0126, 0127, 0128ab, 0142, and 0157 (Murinda and Oliver

2006, (FDA-CFSAN 2007a, Monday et al. 2007). Some infectious strains ofE. coli result in

urinary tract infections (FDA-CFSAN 2007b), meningitis, or other non-diarrheagenic diseases.

However, most pathogenic E. coli cause some type of diarrhea. The primary cause of diarrhea

resulting from E. coli infections is due to release of Shiga toxins (CDC 2008), but not all E. coli

that cause diarrhea possess the Shiga toxin-producing genes. Clinical presentation of diarrhea

can differ greatly among diarrheagenic strains ofE. coli, and among patients, so that grouping of

strains based on symptoms can be ambiguous or misleading. Therefore, laboratory identification

of suspected E. coli bacteria samples should be performed to identify the particular serotype.

Enterovirulent strains are those E. coli that can be categorized by virulence factors (VFs),

which are unique within each group: enterohemorrhagic E. coli (EHEC), enterotoxigenic E. coli

(ETEC), enteropathogenic E. coli (EPEC), enteroinvasive E. coli (EIEC), enteroaggregative E.

coli (EAEC) and diffusely adherent E. coli (DAEC) (FDA-CFSAN 2007b). Of the enterovirulent

E. coli, EHEC causes the most food-borne outbreaks, and the primary serotype is E. coli

0157:H7. Note that additional serogroups 026, 0111, 0126, and 0103 are non-0157

serogroups which have recently resulted in infectious disease outbreaks (Food Source 2006).

However, the focus of this review is limited to EHEC, E. coli 0157:H7, a diarrheagenic and

enterohemorrhagic serotype.

Diarrheagenic strains include those featuring the clinical symptom, namely diarrhea, that

accompanies enteric E. coli infections. However, it should be noted that diarrhea can result from

infections of other pathogens, so that diarrhea is not necessarily indicative ofE. coli infection. In









fact, diarrhea is not even limited to bacterial infections. For example, most diarrhea in children is

caused by two viruses: norovirus and rotavirus (Doyle and Beuchat 2007). Gastrointestinal

diseases, regardless of causative organism, generally always result in diarrhea from a sloughing

off of the epithelial cells lining the intestine.

Enterohemorrhagic E. coli strains are a subset of diarrheagenic strains. These disease-

causing E. coli strains are limited to those that cause bloody diarrhea. Norovirus, rotavirus, and

adenoviruses also can all result in bloody diarrhea (Doyle and Beuchat 2007). Within E. coli

strains, several serotypes are enterohemorrhagic, particularly those that possess the Shiga toxin-

producing genes. Some strains of E. coli overlap into other areas of pathogenicity, because they

possess multiple virulence factors. For example, although E. coli 055:H7 is primarily

enteropathogenic, it is also enterohemorrhagic. Thus, it is often grouped together with 0157:H7

into an "0157:H7 complex" of enterohemorrhagic E. coli strains (Feng and Monday 2005).

Escherichia coli 0157:H7

Within the E. coli 0157:H7 complex, E. coli 0157:H7 is an enterohemorrhagic strain that

possesses many disease-causing virulence factors (Janda and Abbott 2006). This pathogen is

known as Shiga toxin-producing E. coli (STEC) because it produces Shiga toxins, and is also

referred to as enterohemorrhagic E. coli (EHEC) because it is an enteric bacteria that causes

diarrhea and dysentery (Janda and Abbott 2006).

Escherichia coli 0157:H7 and Cattle

Escherichia coli 0157:H7 is a zoonotic bacterium that causes human disease. Cattle are

the primary reservoir (Hussein et al. 2003, Davis et al. 2005, Sargeant et al. 2005, Alam and

Zurek 2006) although other species of domestic animals reared for food also serve as reservoirs,

including sheep (Chapman et al. 1997, Keen et al. 2006), goats (Pao et al. 2005, LeJeune et al.

2006), swine (Chapman et al. 1997, Keen et al. 2006) and poultry (Chapman et al. 1997).









According to a report issued by USDA:APHIS:VS (2007), recent cattle herd prevalence rates in

the United States ranged from 22-100%, although prevalence rates for individual animals within

the herds ranged from 0-9.5%. Hancock et al. (1997) found the prevalence of 0157 lowest in

adult cattle (0.4%) and highest in weaned heifers (1.8%). He also determined that E. coli

0157:H7 of the cattle gut is transient, with fecal shedding lasting a median of 30 d. Infection

with E. coli 0157:H7 is not limited to livestock and poultry as it has also been isolated from

many other mammals and arthropods including deer (Asakura et al. 1998, Dunn et al. 2004a),

opossum (Renter et al. 2004b), pigeons (Morabito et al. 2001), rabbits (Scaife et al. 2006,

Fremaux et al. 2008) house flies (Alam et al. 2004, Sanderson et al. 2006), blow flies (Fotedar et

al. 1992) and slugs (Sproston et al. 2006).

Although adult cattle are asymptomatic carriers ofE. coli 0157:H7 bacteria (Porter et al.

1997), E. coli 0157:H7 can cause disease in unweaned cattle that do not possess a fully

developed rumen, making their digestive system comparable to that of humans. In adult cattle, E.

coli 0157:H7 is a commensal microorganism in the rumen, which helps provide dietary nutrients

to the host animal (Frandson 1969). Caprioli et al. (2005) reported that poultry and pigs do not

serve as EHEC sources; however, other researchers reported contradictory results (Doane et al.

2002). Caprioli et al. (2005) concluded that poultry and pigs become infected by exposure to

cattle and other ruminant (sheep, goats, water buffalo, and deer) excrement.

Although all ruminants may potentially serve as reservoirs for E. coli 0157:H7, cattle

serve as a dominant reservoir ofE. coli 0157:H7, and are considered by many as the primary

reservoir (Loneragen and Brashears 2005). Feedlot cattle pre-harvest diets have been modified in

various ways in an effort to reduce the levels ofE. coli 0157:H7 in feedlot cattle. Addition of

microorganisms to the diet appears to be one of the most promising diet modifications. In









contrast, addition of brown seaweed does not seem to reduce levels of E. coli 0157:H7

(Loneragen and Brashears 2005).

In addition to modification of diet, administration of vaccines, sodium chlorate, and

neomycin sulfate to feedlot cattle has demonstrated beneficial results towards reduced E. coli

0157:H7 pre-harvest levels. In contrast, addition of chlorine to water sources does not appear to

reduce E. coli 0157:H7 levels in pre-harvest feedlot cattle (Loneragen and Brashears 2005).

Escherichia coli 0157:H7 Outbreaks

The origin of the enterohemorrhagic strain of E. coli has not been identified, although it

emerged very recently. It was first identified, although not associated with disease, in 1975 by

the Centers for Disease Control and Prevention (CDC) (Riley et al. 1983). Between 1978 and

1983, laboratory analysis of stool specimens from six sporadic gastrointestinal cases resulted in

isolation of E. coli 0157:H7: five of the cases were for patients with clinical symptoms of

hemolytic colitis, while the sixth was isolated from a patient with an unknown medical history.

In addition to isolation from sporadic cases, E. coli 0157:H7 was isolated and determined to be

the causative agent for three 1982 gastrointestinal outbreaks. The first two occurred in Oregon

and Michigan, USA, and caused 47 known illnesses among fast-food patrons (Riley et al. 1983).

The third outbreak in 1982 resulted in 31 cases, 4 hospitalizations, and one fatality among a

population of 353 elderly persons in Ottawa, Ontario, Canada (CDC 1983). Thus, within a very

short span of seven years, a newly emerged E. coli strain quickly expanded its pathogenic impact

upon the human population.

Denny et al. (2008) and Feder et al. (2003) cite annual E. coli 0157:H7 disease estimates

provided by Mead et al. (1999) for the United States. They estimate that approximately 73,000

cases and 61 fatalities occur each year in approximately 500 outbreaks. An estimated 27-40% of

stricken individuals progress to either severe hemorrhagic uremic syndrome (HUS) or renal









sequelae. They infer that the number of illnesses, hospitalizations and deaths due to E. coli

0157:H7 may be under-reported, due to different reporting mechanisms between states and

among doctors. As of 1999, all food-borne illnesses, including but not limited to E. coli

0157:H7, were determined to be responsible for as many as 5,000 fatalities, with 325,000

hospitalizations out of a total of 76 million illnesses (Mead et al. 1999, Mai et al. 2006). It is

possible that some of the cases of unknown etiology were due to undetected E. coli 0157:H7.

Shiga toxin-producing E. coli were responsible for an estimated 100,000 cases with 2,000

hospitalizations and 91 deaths (Frenzen et al. 2005 as cited in DuPont 2007). For humans,

hospital expenses for patients with infectious diarrhea can be as high as $24,000 versus $9,000

for patients that do not have infectious diarrhea (Suda et al. 2003). Similarly, medication

expenses can be four times higher (-$4,000 vs. -$1,000) and the length of hospitalization can be

three times longer (22 vs. 7 days) (Suda et al. 2003). Economic impacts can also include lost

income for families that must miss work, as well as lost profit for the employers of affected

persons (CDC 2002). Shiga toxin-producing E. coli were responsible for an estimated 100,000

cases with 2,000 hospitalizations and 91 deaths.(Frenzen et al. 2005 as cited in DuPont 2007).

Total numbers ofE. coli 0157:H7 cases worldwide from 1982-2006 reflects movement

into new geographical regions. During this 24-yr period, Doyle et al. (2006) reported worldwide

E. coli 0157:H7 statistics garnered from reviews of the published literature as: 207 outbreaks

and 26,179 cases. The economic impacts of diarrheal disease in the cattle industry are also

significant. Increased operating expenses are observed for dairy farms, cattle rendering plants

and other livestock and poultry businesses that must increase surveillance and management

measures to comply with increased federal regulatory mandates (CDC 2002). Courts can impose

stiff fines upon many different industries including producers, distributors, and restaurants.









Escherichia coli 0157:H7 Pathogenicity

Escherichia coli 0157:H7 possesses multiple virulence factors, including production of

Shiga toxins (Stxl and Stx2) (Renter et al. 2004a), also termed verotoxins (Kruger et al. 2007).

Escherichia coli 0157:H7 is closely related to other STEC organisms, including .\lng//t

species, as well as other strains of E. coli, such as E. coli 055:H7, O111 :H8 and 026:H11

(Renter et al. 2004a). Of the Shiga toxin-producing E. coli strains, E. coli 0157 is isolated from

humans most often, and is responsible for most cases of HUS (Wang and Reeves 1998). Tarr et

al. (1995) reported that E. coli 0157:H7 was responsible for approximately two-thirds of HUS

cases in North America and Europe. The second-most common E. coli serotype that causes HUS

are E. coli 0111 in the United States and E. coli 026 in Europe (Monday et al. 2007). In Europe,

a strain ofE. coli 0157 (SFO157) that, unlike E. coli 0157:H7, does not ferment sorbitol, has

been implicated as a cause of HUS (Monday et al. 2007).

Infections ofE. coli 0157:H7 can lead to severe symptoms including hemorrhagic colitis

(HC), and acute kidney failure due to hemolytic uremic syndrome (HUS) (Ogden et al. 2001).

Hemolytic uremic syndrome is the most common cause of acute kidney failure in North America

(Karmali 1989). In the United States, HUS is the primary cause of kidney failure in children

(Breuer et al. 2001). Since it was initially identified in 1982 (Riley et al. 1983), E. coli 0157:H7

has caused enteric outbreaks with 8% (Slutsker et al. 1998) and 9% (Bell et al. 1994, CDC 2009)

of cases progressing to HUS (Slutsker et al. 1998). Progression of E. coli 0157:H7 infections to

HUS occurs throughout the world, especially in developed countries (Ooka et al. 2009). In Japan,

6% of persons infected with E. coli 0157:H7 developed HUS (Watanabe et al. 1999). In

Scotland, E. coli 0157:H7 has been determined responsible for more than 90% of HUS cases

(Pollack 2005).









In the United States, infection with E. coli 0157:H7 was made a nationally notifiable

disease in 1995 (CSTE 2005). Enterohemorrhagic E. coli 0157:H7 (EHEC) is commonly found

in association with livestock, especially cattle (Rasmussen et al. 1999, Hussein and Sakuma

2005). In the United States, bacterial outbreaks due to E. coli 0157:H7 have occurred with

increasing frequency since the discovery of this pathogenic strain (Hancock et al. 1994).

Escherichia coli 0157:H7 Prevalence and Persistence

Escherichia coli 0157:H7 can persist in a wide variety of substrates for varying lengths

of time. These substrates include livestock manure, human waste, garbage, compost, soil, and

food crop plants, animal watering troughs, and natural water sources such as lakes, rivers and

puddles (Avery et al. 2008). Persistence of E. coli in livestock fecal matter has been observed in

manure from dairy cattle, beef cattle, swine, and sheep. Escherichia coli 0157:H7 has also been

observed to persist for up to three days in the acidic gut and feces of the house fly (Kobayashi et

al. 1999).

Escherichia coli 0157:H7 can survive for up to 2 yr in dairy farm environments (Shere et

al. 1998). This increases its potential for disease transmission. Longitudinal prevalence (i.e.,

persistence over long period of time) of E. coli on dairy farms is seasonal, with highest

prevalence in spring and late summer (Hancock et al. 1997, shere et al. 1998, Vidovic and

Korber 2006). Persistence of E. coli 0157:H7 has been observed in soil (Islam et al. 2004), water

(Sargeant et al. 2003) and for up to 21 mo in composting manure (Kudva et al. 1998). Islam et al.

(2004) observed that E. coli 0157:H7 can survive in soils amended with cattle manure for more

than five months.

Persistence of E. coli 0157 and specifically, E. coli 0157:H7, in farm environments has

been linked to its prevalence in water, including water tanks (Sargeant et al. 2003) and open

water (Shere et al. 1998). Sargeant et al. (2003) reported higher individual cattle shedding rates









for beef cattle in feedlot pens that tested positive for E. coli 0157 than for beef cattle in pens that

tested negative for E. coli 0157. They also observed that E. coli 0157 was more likely to infect

the water if it was detected in water-tank sediment.

Our understanding of the ability of E. coli 0157:H7 to persist on plants has been further

understood by recent evidence that plants take up the bacterium through their stomata and

translocate the organism to their tissues (Teplitski et al. 2009) However, isolation frequency of

E. coli 0157:H7 serotype has been lower than that of other enteric pathogens (Tyler and Triplett

2008). The ability to find "safe harborage" inside plant tissues could potentially permit greater

persistence in plants than is possible by mere surface contamination. Persistence ofE. coli

0157:H7 and other enteric pathogens in and on plants is closely associated to the common

agricultural application of animal manures (Brandl 2006).

Escherichia coli 0157:H7 Detection, Isolation and Identification

Several different selective media have been used to detect, isolate and identify E. coli

057:H7 in samples collected from dairy cattle and other livestock farm environments, including

Sorbitol MacConkey (SMAC) agar, Levine's Eosine-Methylene Blue (L-EMB) agar, and

CHROMAgar. These media contain selective agents, such as dyes and bile salts that inhibit the

growth of competing microbial organisms, such as gram positive bacteria. Their specificity is

increased further by addition of antibiotics (Heuvelink 2002). Although E. coli 057:H7 is

difficult to isolate from fecal samples, it occurs in low densities as compared to other enteric

microorganisms. In fecal samples, enrichment culture provides more sensitive isolation ofE. coli

0157:H7 than direct culture (Sanderson et al. 1995, Zhao et al. 1995, Wallace and Jones 1996),

largely due to the inclusion of selective agents and/or antibiotics.

Although selective media inhibit competing bacterial growth to improve recovery of E.

coli 0157:H7, each selective medium used to isolate and identify E. coli 0157:H7 has









disadvantages. For example, although SMAC agar contains selective agents to increase its

specificity for isolation and identification ofE. coli 0157:H7, SMAC agar has poor overall

specificity. Another disadvantage of SMAC agar is its propensity to change colors when

incubated for prolonged periods, making it difficult to interpret results.

Although supplementation of microbiological media, both selective and non-selective

(general growth) types, with cefixime and potassium tellurite increases specificity, such

supplementation can result in false positives, particularly when genetically similar enteric

bacteria such as P. mirabilis or E. hermanii (Wallace and Jones 1996) are competing for growth.

In addition to providing false negative results, supplementation with antibiotics and/or bile salts

can actually inhibit recovery of acid- or freeze-stressed E. coli 0157:H7, regardless of whether

the media used is selective or non-selective (Stephens and Joynson 1998).

Escherichia coli is commonly found in complex media such as manure (Pell 1997).

Although widely present in fecal matter, E. coli 0157:H7 is difficult to isolate, because there are

also high concentrations of competing, "background" microorganisms (BM), such as Proteus,

Klebsiella, Salmonella, and .\nge//ll (Vold et al. 2000). Escherichia coli populations decrease

rapidly after host excretion, because of changes in temperature, light, pH and moisture conditions

(Unc and Goss 2006), and from increased growth rates of competing bacteria. Historically, the

largest concentrations of E. coli have been obtained from fresh manure, either during or

immediately after deposition.(Bolton et al. 1999, Duffy 2003). More recently, still higher

numbers of E. coli have been collected by insertion of anal-rectal swabs (Chapman et al. 1997,

Pearce et al. 2004, Ahmad et al. 2007), a sampling method that is considered by some to be more

sensitive than collection of deposited manure (Rice et al. 2003). However, Vidovic and Korber









(2006) determined that large 10-g fecal samples provided more accurate isolation of E. coli than

swabs.

Isolation of E. coli strains from manure is complicated by the large numbers and variety

of different bacteria species present. Kudva et al. (1998) and Tutenal et al. (2003) observed that

animals inoculated with 105 CFU/g of E. coli had average initial E. coli concentrations of 105 -

108 CFU/g in fresh feces, that concentrations in manure piles remained at 105 106 CFU/g for

approximately one year, but decreased to 101 102 CFU/g when aerated (Tutenel et al. 2003,

Dunn et al. 2004b) determined that prevalence rates of E. coli 0157 in dairy cattle manure were

commonly < 1%, and never exceeded 5%.

Pearce et al. (2004) tested up to three samples from individual cow pats, and determined

that E. coli 0157 distribution within single cow pats is highly variable; thus, prevalence rates can

vary significantly, depending on where in the cow pat samples are obtained. Echeverry et al.

(2005) also tested multiple samples per cow pat, and determined that prevalence rates increased

from 8.2% when one sample/pat was tested, and up to 20% when 5 samples/pat were tested.

Isolation of E. coli strains differs for diarrheagenic versus non-diarrheagenic species

(Heuvelink 2003). Within diarrheagenic E. coli strains, differentiation of enterohemorrhagic E.

coli (EHEC) 0157:H7 can be accomplished by any of several methods, and there is no general

consensus on which method is superior. Typical E. coli biochemical characteristics include the

following: Lysine +, Citrate -, Indole +, Acetate +, Lactose +, and aerogenic + (production of gas

with carbohydrate metabolism).

Escherichia coli 0157:H7 does not ferment sorbitol within 24 h (Doyle and Schoeni

1984), and is therefore described as "non-sorbitol-fermenting" (Desmarchelier et al. 1998).

Because most isolation culture methods involve 24-h incubation, this characteristic provides easy









differentiation from similar and closely related enteric bacteria species. Escherichia coli

0157:H7 has a characteristic morphology and color when grown on Sorbitol MacConkey agar

(SMAC) supplemented with selective antibiotics such as potassium tellurite and cefixime (CT-

SMAC) that inhibit gram-positive bacterial growth and enhance gram-negative enteric bacterial

growth. Such media is commonly used, although the percentages of the respective antibiotics

may differ among researchers. Typically, antibiotic concentrations are: potassium tellurite (1.25

[tg/l), sometimes referred to simply as "tellurite," (Alam and Zurek 2004) and cefixime (15 [tg/l)

(Desmarchelier et al. 1998). Antibiotics are added to provide selective growth of enteric bacteria,

especially of E. coli 0157:H7, as well as to provide differentiation of E. coli 0157:H7 from its

close enteric relatives. When cefixime and potassium tellurite are supplemented into SMAC, the

resulting agar is referred to variously as CT-SMAC, SMAC-CT, or even as mSMAC, where the

"m" represents "modified," even though the type of modification may differ dramatically.

Comparison of results obtained from use of SMAC modified with antibiotics needs to be

conducted very carefully, for upon examination, not all researchers use the same concentrations

of antibiotics. Thus, while multiple researchers may all refer to "CT-SMAC," they may be using

fundamentally different selective media. Additionally, some researchers add yet another

antibiotic, vancomycin; again, with differing concentrations. In some rare instances, the

antibiotic cefsulodin has been used, either in addition to those listed previously, or as a

replacement for one or more of them. Cefsulodin is recommended along with cefixime and

potassium tellurite in an FDA protocol for isolation ofE. coli 0157:H7 (FDA-CFSAN 2007b).

There is one caveat regarding the use of potassium-tellurite antibiotics in media.

Although primarily used to select for non-sorbitol fermenting E. coli 0157:H7, potassium

tellurite also permits growth of other Shiga toxin-producing E. coli strains such as 026, 0111,









and 0145 which possess the ter gene that confers resistance to this antibiotic. Possession of the

ter gene appears to be positively correlated with possession of the eae gene. Therefore, use of

this antibiotic in selective plating media makes isolation of 0157:H7 more likely, but not

guaranteed. Strains of E. coli isolated with potassium-tellurite tend to possess the eae gene (Orth

et al. 2007).

Even without supplemental antibiotics, SMAC is selective for gram-negative bacterial

growth, because crystal violet is a selective component of sorbitol MacConkey agar (SMAC) that

inhibits growth of competing enteric gram-positive bacteria (Doyle and Beuchat 2007). For this

reason, SMAC is widely considered to be the best media to use for successful isolation ofE. coli

0157:H7 (CDC 1994). Chou et al. (2000) observed that E. coli 0157:H7 cultured in non-

selective general-growth trypticase soy broth (TSB) culture tubes for one day at the low-

temperature stresses of -5 C, -18 C and -28 C exhibited respective survival rates of 87.55%,

0.72%, and 1.66%, when followed with a subsequent 1-h exposure to crystal violet. This high

mortality of cold-stressed E. coli 0157:H7 after 1 h exposure to crystal violet illustrates the

importance of crystal violet dye for the selective isolation of E. coli 0157:H7, even without

antibiotics. The use of antibiotics is helpful against competing gram-negative bacteria after

crystal-violet inhibition of gram-positive bacteria.

Although it may seem frustrating that researchers use various concentrations and

mixtures of antibiotics to supplement SMAC for selective growth and isolation of E. coli

0157:H7, there is ample justification for it: different strains or serotypes of E. coli 0157:H7 may

react dissimilarly when exposed to environmental stresses. Development of antibiotic resistance

in a given bacterial population may change an organism's ability to grow in antibiotic-treated

media. Escherichia coli 0157:H7 is typically resistant to many antibiotics, and will usually grow









on media that contains cefixime and potassium tellurite, whereas other enteric bacteria will be

inhibited by these two antibiotics (Desmarchelier et al. 1998).

While CT-SMAC, in all its variations, appears to be the most selective and most effective

media for differential isolation ofE. coli 0157:H7, direct plating may not be sensitive enough to

identify the low levels of 157:H7 that are typical of samples with low concentrations of E. coli

0157:H7 such as exist in many food samples (Willshaw et al. 1994). This might also be true for

fecal samples, particularly because cattle shed E. coli 0157 both seasonally and sporadically, so

that bacterial concentrations may vary widely from one sampling occasion to the next (Matthews

et al. 2005). This lack of sensitivity led to the development and wide use of immunomagnetic

separation (IMS) for selective isolation ofE. coli 0157:H7 on magnetic beads coated with anti-

E. coli 0157:H7 antibodies. IMS permits very selective binding of E. coli 0157:H7 to the beads.

The conjugated bacteria-bead complex is manually separated from bacterial enrichment broths

magnetically and by simultaneous rinses in buffered detergent, such as PBS Tween-20. The

benefit of IMS is that it permits a more sensitive detection of E. coli 0157:H7 than traditional

direct plating (Grif et al 1998). However, the disadvantage of IMS is that specificity may be

decreased by the many sorbitol non-fermenting microorganisms that bind non-specifically to the

immunomagnetic beads (Chapman and Siddons 1996).

Standard microbiological selectivity of isolation has been greatly improved by

immunomagnetic separation (IMS) technology which increases sensitivity 100-fold over direct

culture methods (Karch et al. 1996). Immunomagnetic separation utilizes magnetic beads coated

with polyclonal pathogen-specific antibodies into specimen enrichment samples for inoculation.

The magnetic beads specifically bind to the 0157 somatic antigen of E. coli 0157. Sequential

rinsing steps remove any sample debris and/or non-specifically bound pathogens from the beads,









and the beads are then plated directly onto selective media (Heuvelink 2003). The IMS protocol

is especially effective in samples such as feces which contain large populations of competing

(background) microorganisms (Manafi 2003).

Detection ofE. coli 0157:H7 by direct culture is an expensive, difficult, labor-intensive

and time-consuming process with little consistency in chosen isolation methods among

researchers (Manafi 2003, Heuvelink 2003). Standard microbiological tests typically take 3-4

days to complete and require repetitive re-plating of suspect organisms onto new media each

day. High background microfauna concentrations typical of fecal samples greatly increase the

difficulty with which E. coli 0157:H7 is successfully isolated. Competing microorganisms can

be so numerous that they result in lawns of bacterial overgrowth, even when plated onto selective

media plates that contain antibiotics specifically designed to inhibit the growth of non-E. coli

0157:H7 organisms (Heuvelink 2003). Identification by biochemical and morphological traits

can take as long as 7 d, depending on how many traits are examined and which selective media

are used. Biochemical tests have low specificity and selectivity (Visetsripong et al. 2007),

making it difficult to accurately identify and isolate 0157:H7, especially if present in very low

numbers. Successful detection and isolation of E. coli 0157:H7 is further confounded by the

presence of closely related bacteria species. This is particularly evident in fecal environments

where competing microorganisms share similar phenotypical and biochemical traits with E. coli

0157:H7, even when isolates are plated on selective media designed to differentiate between

bacteria species. Thus, isolation of individual colonies with typical E. coli 0157:H7

characteristics on any particular media require subsequent transfer to additional selective media

to confirm identification.









Rapid, selective and sensitive detection methods are important (Ogden et al. 2001).

Although addition of inhibitors and/or antibiotics to growth media increases the selectivity of the

media for successful isolation of the target bacteria, it often decreases the sensitivity, so detection

of small numbers of bacteria becomes increasingly more difficult.

Escherichia coli 0157:H7 and DNA-based Isolation Techniques

Polymerase chain reaction (PCR) provides a more selective and sensitive method than

direct-culture to identify E. coli 0157:H7 in samples (Visetsripong et al. 2007), and permits

serotyping (Beutin et al. 2007). PCR can be performed in 6 hours (Szalanski et al. 2004),

making it more promising for timely identification ofE. coli 0157:H7. Primer pairs can be used

individually in uniplex or combined in multiplex PCR to amplify one or more specific target

gene fragments, respectively.

Shiga toxin (Stx) gene fragments are often targeted in PCR because they are probably the

main virulence factors (VFs) ofE. coli 015:H7 that lead to HUS and HC (Beutin et al. 2007,

Kruger et al 2007). Successful PCR amplification of Stx gene fragments confirms the presence of

the Shiga toxin-producing gene, but does not specifically confirm E. coli 0157:H7, because Stx

genes are also present in more than 200 serotypes of E. coli (Beutin et al. 2007) and in closely

related .\ligel,// spp. (Donnenberg 2002).

While PCR provides quick identification of positive E. coli 0157:H7 samples, one

disadvantage of using PCR is that fecal components inhibit polymerase chain reaction, and thus

result in false-negative PCR results (Wilde et al. 1990, van Zwet et al. 1994). However, these

chemicals can be removed by detergents (Wilde et al. 1990). Another disadvantage of PCR is

that there is evidence that E. coli 0157:H7 strains can lose Shiga toxin-producing genes (Feng et

al. 2001) so that PCR might not detect positive E. coli 0157:H7 samples that are detected by

serological or biochemical methods.









Summary

While cattle are a primary reservoir of E. coli 0157:H7, they shed this pathogen

intermittently in their feces (Wetzel and LeJeune 2006). The highest cattle shedding ofE. coli

0157:H7 occurs during warm summer months when both E. coli 0157:H7 and house fly

populations are also at their highest levels (Alam and Zurek 2004). Standard fecal cultures based

on 25 g of cattle feces inoculated in 225 ml of a non-selective nutrient broth typically detect E.

coli 0157:H7 concentrations that range from 2 x 102 to 8.7 x 104 CFU/ml. (Hancock et al.

1997). In contrast, adult house flies weighing only 0.13 g have E. coli 0157:H7 in concentrations

ranging from 3 x 101 to 3.0 x 105 CFU/fly (Alam and Zurek 2004), the equivalent of 3 x 102 to

3.0 x 106 CFU/ml, so there is as much as a 100-fold higher concentration ofE. coli 0157:H7 in

house flies than in manure. Thus, it is possible that E. coli 0157:H7 on dairy farms might be

more accurately detected by testing adult house flies instead of cattle manure samples, regardless

of which isolation method is utilized.









CHAPTER 2
HOUSE FLY DISPERSAL

Introduction

Musca domestic L., the house fly, is capable of transmitting more than 100

pathogenic organisms (Greenberg 1973) that can cause disease in humans. Musca

domestic has especially strong links to enteric diseases, such as typhoid, cholera,

dysentery, and diarrhea (Steinhaus 1946), which cause high mortality rates worldwide,

particularly in children of developing countries (Kosek et al. 2003, iOWH 2008). The

house fly's ability to transmit pathogens is due to the following synergistic factors: 1) its

predilection to breed in fecal or rotting organic material that may be teeming with

disease-causing microorganisms, 2) its habit of constant regurgitation and excretion while

eating, and 3) its ability to disperse over wide geographic areas as far as 33 km (20 mi)

(Murvosh and Thaggard 1966, Meerburg et al. 2007), including direct flight over large

swamps and across rivers 300-500 m wide (Shura-Bura et al. 1962). Considered alone,

none of these behaviors creates a human disease threat. However, considered together,

the potential for disease transmission expands exponentially, as the house fly can

potentially transmit many fecal pathogens to any food source within its dispersal range.

Dispersal of the pathogen-infected house fly from a dairy to a town introduces and

increases the potential for disease transmission to humans.

Flight and dispersal behavior for house flies and other species of synanthropic

flies with similar breeding habits, such as stable flies and blow flies, has been reported by

a number of researchers and varies drastically in individual studies. In rural areas, house

flies can disperse 12 km (Broce 1993a) and have been documented dispersing up to 21

km (Bishopp and Laake 1921, Alam and Zurek 2004) from their breeding sites. In urban









communities, most flies disperse within 1.7 km (1 mi) of release sites (West 1951,

Quarterman et al. 1954, Schoof and Siverly 1954b; Hanec 1956; Sacca 1964, Milio et al.

1988). However, house flies have been documented to disperse distances up to 33 km in

urban environments (Murvosh and Thaggard 1966). House fly dispersal speed has been

documented at a rate of 1 km/h for the first 3-4 h, when dispersal occurred as direct flight

over a large swampy area and across rivers (Shura-Bura et al. 1962). Dispersal distances

and recapture rates might be influenced by the type of flies used, i.e., field-collected or

laboratory-reared. Previous studies have indicated that use of field-collected flies is more

representative of dispersal under natural conditions than flies that are reared for multiple-

generations in the laboratory. Eddy et al. (1962) recaptured a 10-fold higher percentage

of field-collected flies than laboratory-reared flies, implying that laboratory colonies may

lose their ability to disperse.

Dispersal of house flies increases the potential for transmission of zoonotic

diseases to humans, particularly from sites conducive to fly breeding such as dairies that

house large numbers of cattle (Kaufman et al. 2005b, Ahmad et al. 2007, Conn et al.

2007), beef cattle feedlots (Skoda et al. 1993, Thomas 1993, Sanderson et al. 2006),

swine facilities (Rosef and Kapperud 1983, Halverson 2000), poultry buildings (Hald et

al. 2004, Watson et al. 2007) as well as non-agricultural sources of fecal waste, such as

dog excrement (Wilton 1963), to nearby human population centers. If house flies can

maintain a travel speed of 1 km/h for an extended period of time, and if house fly

dispersal flight occurs in a straight line from a breeding site, then house flies could

potentially transmit infectious pathogens as far as 12 km in only 12 h (Meerburg et al.

2007).









The house fly readily alights on decomposing fecal/organic matter as well as

human food, and moves freely between the two. External contamination of house flies

can range from 2.5 to 29.5 million bacteria per fly (Hawley et al. 1951), and some

bacteria can survive up to 3.5 d on the surface of house flies (Peppler 1944). Bacterial

contamination of house flies can also occur after flies contact food crops that have been

fertilized with liquid manure or solid fecal waste (Islam et al. 2005). Mechanical

transmission of this bacterium by house flies has been well-established by many

researchers (Echeverria et al. 1983, Fotedar et al. 1992, Sasaki et al. 2000, Alam and

Zurek 2004, Buma et al. 2004, Ahmad et al. 2007, Nmorsi et al. 2007). Biological

transmission also appears likely if E. coli 0157:H7 is capable of replicating within the

house fly gut (Hawley et al. 1951, Petridis et al. 2006).

House fly dispersal has been measured by many types of mark, release and

recapture studies using fluorescent dusts, sticky traps, and UV lights (Hogsette 1983,

Osek 2001). One of the easiest and most efficient techniques for marking and releasing of

large numbers of small insects is the application of fluorescent dust (Hagler and Jackson

2001). Insects such as house flies are collected in the field or mass-reared in the

laboratory, marked for future identification, released in the field, and recaptured at

various distances from their release site. Ideally, the substance used to mark the insects is

long-lasting, is non-toxic to both the insect and the environment, does not change insect

behavior, and is easy to observe on recaptured specimens. Additionally, the ideal insect

marker is inexpensive, readily available, has a long shelf life, performs consistently, and

can be applied quickly and easily (Osek 2001). Quick and easy application of markers is









particularly beneficial in the field, where access to tools used to decrease insect activity

for easier application of markers, such as chill tables or CO2 anesthetic, is limited.

Many types of markers have been used effectively in the past, but are no longer

recommended or are sometimes prohibited under existing regulatory legislature, due to

human, animal and/or environmental health concerns. For example, radioactive

phosphorus (32p) has been added to adult fly laboratory diets and 32P-labeled flies

released and recaptured in the field (Lindquist et al. 1951, Yates et al. 1952, Eddy 1962,

Shura-Bura et al. 1962, Williams, J. R. P. 1973). Marked flies were subsequently counted

using readings on Geiger-counters (Hoffman and Lindquist 1951; Lindquist et al. 1951;

Yates et al. 1952; Eddy 1962; Shura-Bura et al. 1962; Williams, J. R. P. 1973). Hoffman

and Lindquist (1951) reared flies in media containing 32P to compare the efficacy of this

method against application by ingestion, and determined that feeding 32P to adult flies

was both more effective and cost-effective. Lindquist et al. (1951) compared marking

adult house flies by adding 32P to the diet against dusting with fluorescent dusts. They

determined that marking with dietary 32P was more efficient and less labor-intensive, and

they observed that the dusts wore off within 48 h so that identification of dusted flies was

difficult. Similarly, Eddy et al. (1962) concluded that ingestion of 32P was a more useful

marking method than fluorescent dusts, because wild flies had natural fluorescence that

was easily confused with the fluorescent marker (C-205 yellow, Ultra Violet Products,

Inc.) used in their study.

Although there is no universal method of marking insects, dusts are possibly the

most frequently used external markers, due to their ease of use in both application and

observation, as well as their low cost, ready availability, and low toxicity. Fluorescent









dusts, in particular, Day-Glo powdered pigment dusts, have been used to track dispersal

and population dynamics, without any observed adverse changes to insect behavior or

mortality (Hogsette 1983, Hogsette 1984, Kristiansen and Skovmand 1985). Fluorescent

dusts also offer potentially long-term investigative study possibilities, because the dust

has been shown to last up to 3.5 mo in the field (Taft and Agee 1962). Flies are dusted

with fluorescent powder, released, and recaptured; subsequent UV light examination of

recaptured flies illuminates any retained fluorescent dust on areas of the body that the fly

has difficulty grooming. Flies dusted as adults will typically retain dust particles on

portions of the thorax; when the puparia are dusted, flies emerge, crawl through the dust,

and retain it on their ptinilum (Hogsette 1983).

An additional advantage of fluorescent dusts is that their visibility is greatly

enhanced when examined under long-wave ultraviolet (UV) light. Thus, large numbers of

recaptured house flies on sticky traps can be examined rapidly and easily under UV light

to determine how many are marked. This eliminates time- and labor-intensive

observation methods used with alternative marking techniques, as there is no need to

destroy individual insects to observe internally expressed dyes, to apply solvents, or to

perform genetic analysis. Application of fluorescent dusts is relatively easy, inexpensive,

and less labor-intensive than other insect marking techniques, and enables marking of

thousands of insects simultaneously (Zhao et al. 1999). Additionally, application of

fluorescent dusts can be accomplished using mechanical dusters (Hogsette et al. 1993).

Flies can be captured on a wide variety of traps. Alsynite traps have proven

effective, easy to transport, and easy to use in the field. One type of alsynite trap consists

of a translucent rectangular fiberglass panel folded to form a cylinder that is then









wrapped with an adhesive coated clear plastic sleeve (Hogsette and Ruff 1990). The

fiberglass panels reflect ultraviolet light, making them attractive to many flying insects

such as stable flies, Stomoxys calcitrans (L.), (Williams, D. F. 1973), alate red imported

fire ants, Solenopsis invicta Buren, (Milio et al. 1988), and house flies (Geden 2006).

Flies that land on the adhesive become stuck. Traps can be examined under UV light to

observe fluorescence on individual flies, and flies can be identified to species.

Although flies ostensibly have all their physiological needs met in the dairy

environment and have no discernible reason to leave the dairy, there is evidence that they

disperse in all directions without aid of vehicle transport, in direct flight, away from the

dairy. The true pathogen transmission potential ofM. domestic in north-central Florida

can be better estimated if house fly dispersal behavior can be determined. The goals of

this study are: 1) determine if house flies disperse from a dairy to a town, 2) determine

recovery rates of marked flies, and 3) evaluate possible deleterious impact of fluorescent

dusts used to mark the flies.

Materials and Methods

Laboratory Facilities and Rearing

Unless otherwise stated, all laboratory studies and fly rearing were done in the

fly-rearing laboratory at the USDA-ARS-CMAVE in Gainesville, FL. Standard USDA

rearing conditions for all fly stages in the fly rearing chambers were 26 2 C, 60 5%

RH, 12:12 L:D. (Hogsette 1996). Any mention of fly larval growth medium refers to the

Gainesville House Fly Diet (GHFD) (wheat bran 50%, alfalfa meal 30%, cracked corn

20%) (Hogsette 1992), unless another medium is specifically described. Larval growth

medium was moistened with water in a 1:1 (v:v) diet:water ratio. Adult house flies were









provided with food comprised of 6:6:1 powdered milk: granulated sugar: powdered egg

yolk, and ad libitum access to water (Hogsette et al. 2002).

During this study, dairy-collected adult and immature house flies were frequently

reared in the laboratory. Fly transport cages measured 30.5 x 30.5 x 30.5 cm (Model

1450B, Bioquip, Rancho Dominguez, CA) and were used to move adult dairy-collected

flies back and forth between the field and laboratory, and to release marked adult flies at

the field site. Rearing cages were either: 46 x 46 x 46 cm (Model 1450C, Bioquip,

Rancho Dominguez, CA), or 45 x 36.25 x 36.25 cm (USDA) (USDA-ARS-CMAVE,

Gainesville, FL).

Regardless of which cage was in use, adult house flies were maintained under

standard USDA rearing conditions as previously described. Approximately 100 g of fresh

GHFD was placed in each cage in 237-ml Styrofoam deli cups (Model # 8SJ32, Dart

Water Corp., Mason, MI). Approximately 400 ml of water was placed in 473-ml clear

plastic deli cups (Model #L2516, Newspring Packaging, Keamy, NJ), and covered with a

single layer of foam packing pellets to prevent drowning. In adult rearing cages, fresh dry

diet was added to cages every 3 d, and additional containers of water were added if water

levels dropped to less than 25% or if dry diet or water showed mold growth. Transport

cages were in use for only a few hours, so replenishment of dry diet and water was not

necessary.

Description of Study Area

The study area used from 16 October 2008 to 4 December 2008 and from 14 May

to 25 June 2009 is located in north-central Florida, in a geographical region dominated by

small livestock farms, primarily dairies and poultry broiler farms. The area was relatively

flat, with gentle rises (hillocks). Patchy habitats of scrub forest were interspersed









throughout the area, with fields often separated by tree-lines (wind-breaks). Dairy

pastures on the release farm were irrigated with water and fertilized with slurry (liquefied

manure), both of which were applied with water sprayers of various types. The study area

was the area between a small town and the dairy upon which the flies were released. This

dairy is hereafter referred to as "Dairy A." There were two other dairies nearby: both

were adjacent to Dairy A with shared property lines. "Dairy B" was located W of Dairy

A, and "Dairy C" was located S of Dairy A. The town was located entirely within 3.5 km

SSW of Dairy A, so that traps placed at the closest edge of town relative to Dairy A were

located approximately 2.5 km from the dairy, and the trap that was placed in town

furthest from Dairy A (trap 12, at Restaurant D) was located 3.5 km from the Dairy A

release site. House flies were collected from both Dairy A and Dairy B; however marked

adult flies were released only from Dairy A.

Dairy A milked between 400-500 cows and maintained 4-5 bulls at the beginning

of the study. Approximately 50% of the cattle were sold during the summer of 2009, so

the herd decreased to 200-300 by the end of the study. The cattle grazed on Bahia grass

fields during the day, and came to the open-sided barn twice daily to eat grain and be

milked. Hay was provided in the fields to supplement grazing. The barn contained 2-3

large cement water-troughs located mid-way between the south and north barn edges, and

was equipped with misters that sprayed water over the cows, and large fans that increased

air-flow within the barn. The barn was located atop a small hillock, and consisted of a

large cement floor that was slightly graded to enhance twice-daily rinsing of feces and

urine into a cement culvert that emptied into a nearby cement waste-containment system.

The floor was rinsed by flooding it with water that was stored in a large tank located at









the eastern end of the barn at the point of highest elevation. After cattle feeding was

completed, a release valve on the tank was opened, so that the entire contents of the water

tank flooded the floor of the barn, and rinsed fecal waste into the culvert.

The barn had two cement feeding troughs, one along each of the south and north

lengths. The feed troughs were located at ground level, at the edge of the barn's roof drip

line, so that the grain was largely protected from rainfall except during windy conditions.

The troughs did not receive water spray from the barn misters. After rain, large puddles

formed next to the feeding troughs, and some runoff entered the open ends of the feeding

troughs. Grain was poured into the troughs mechanically by a truck twice daily. Cattle

entered the barn approximately 30-60 min before the grain was delivered, and also were

observed eating grain that remained in the troughs from previous feedings, even if it was

slightly moist from rain or infested with fly larvae. The feeding troughs consistently

served as house fly breeding sites during this study. Intermittent applications of toxic fly

bait containing imidacloprid (QuickBaytTM, Bayer, Shawnee Mission, KS) and

insecticides containing permethrin both inside and outside the barn were used to suppress

adult house fly populations. Additionally, permethrin-impregnated ear tags were used for

horn fly, Haematobia irritans (L.), control.

Dairy B was a dairy with 500-600 milking cows, approximately 30 bull calves,

and 7-8 bulls. The cattle spent the majority of their time in open-sided barns, where feed

was available ad libidum. When in the barn, cattle were packed so tightly that they were

pressed together. Cattle were milked twice daily. Cattle sometimes grazed in the grass

fields, and supplemental hay was available in the fields. There were two barns,

respectively placed north and south of the milking parlor, which were identical in









structure. Each barn contained one large cement water trough located in a corner. Several

large ceiling fans were evenly distributed throughout the barns. Water misters were also

located throughout the barns. The barns were located on an inclined patch of land so that

the north barn had the higher elevation. The cement barn floors had little or no grade,

limiting runoff Fecal and urine waste products accumulated rapidly, and were removed

by spraying water with a hand-held hose. Liquefied waste flowed into a nearby earthen

lagoon system situated south and downhill of the south barn.

A large cement feed trough ran east to west through the center of each barn. Feed

was provided daily to the trough by a mechanical feed auger system. The feed troughs

were elevated, so that cattle could consume feed without stooping. Fecal and urine waste

did not appear to contact grains. New feed was apparently added to old feed as troughs

were never completely empty. The troughs were within range of the water misters, and

the feed was frequently damp. The feed troughs were surrounded with steel poles that

provided abundant horizontal and vertical resting surfaces for adult house flies.

Additionally, rough wood support beams were located at regular intervals along the feed

troughs. The barn's exterior was constructed of rough wood, with wood beams extending

from floor to ceiling, and plywood panels covering the top half of the barn. Cattle were

restrained in the barn by a wood fence that surrounded the perimeter of the barn.

The south barn was built so that its southern edge was above-ground. Fecal waste

draining from the south barn usually overflowed onto the ground, and possibly

contaminated spilled grains that were located underneath the auger used to supply new

feed to the feed trough. Overflow of fecal waste into spilled grains was further facilitated

by a constantly flowing hose that supplied the water trough inside the barn. The trough









overflowed out of the south barn onto the ground through which the fecal waste moved

en route to the lagoons. The spilled grains were located slightly downhill of the fecal-

waste ditch. Therefore, grains that spilled onto the cement pad beneath the auger were

moistened by various sources, including rainfall and diluted fecal waste. Between the

south barn and the lagoons, there was a large patch of untended grass and herbaceous

growth.

Some of the calves were kept in individual calf hutches, while others were

permitted free range in a field. This field was not mown during the 2009 6-wk study and

the forage growth was taller than calf-height by the end of the study. The calves were all

located in a pasture approximately 0.5 km south of the feed barns. Calves in hutches were

supplied with feed and water. Hutches were placed in the shade under large deciduous

trees. Feed was provided to the calves in a large plastic drums with and without

protective coverings. Water was provided by insertion of a water hose into a large plastic

drum set on its end.

Similar to Dairy A, fly control was attempted by scattering imidacloprid fly bait

and spraying permethrin around barns. Additionally, permethrin-impregnated ear tags

were attached to cattle ears.

House Fly Collection and Rearing

Adults and immature stages of house flies were collected from both dairies

weekly for this mark, release and recapture dispersal study. Adult house flies were

collected via repeated sweep-netting above the feed troughs for up to 30 min by 1-2

persons, and transferred into a transport cage that already contained water and adult

house fly diet, as previously described. Transfer of flies from sweep nets into transport

cages was accomplished by inverting the net into the cage through a cotton sleeve









attached to the front of the cage, and shaking flies loose inside the cage. Adult flies that

were captured and subsequently released on-site at Dairy A in this manner are referred to

as the sweep netted flies (SN).

The total number of adult house flies captured each week was estimated using the

following two methods. During the first week, an average sweep net count rate was

established for each individual, by having each person collect two extra sweep net

collections, one at the beginning of sweep-netting, and one at the end of sweep-netting.

When dairy-collected house fly populations at the dairy were visually low, the number of

flies caught in initial sweeps was much greater than the number caught in final sweeps.

The first and final sweeps were used to determine average sweep net values per

individual researcher, to compensate for the decreased house fly population in the

vicinity resulting from the sweep-netting impact. After determining each researcher's

average sweep count rate, the total number of house flies collected per individual was

estimated by multiplying the number of sweeps that each person performed by that

individual's respective average sweep net count rate. Finally, the total count of house flies

captured for the mark and release technique was calculated by adding individual counts

together. This method was used only during the first week, because fly numbers appeared

to be reduced too much by this method. Therefore, during the remainder of the study

period, visual estimates of the reared adult house flies were made.

To supplement dairy-collected-caught adult flies, immature dairy-collected flies

were reared in the laboratory (IR). House fly eggs, larvae, and/or pupae were collected by

filling 3.79 liter plastic buckets (Model # 8671-2 NRC 65 MIL, Letica Corporation,

Rochester, MI) with decomposing grains and other dairy sources (dairy-collected









materials) where immature flies were observed prior to collection. Dairy-collected

materials and immature flies were transported back to the USDA laboratory in an air-

conditioned vehicle.

In the laboratory, dairy-collected materials containing IR developing flies of all

immature stages were evenly dispersed among several enamel-coated steel dental pans

(19 x 31 x 5 cm) so that each pan was filled to a depth of approximately 2.5 cm. The

remaining depth of each pan was filled to the top by addition of moistened Gainesville

house fly diet (GHFD) (Hogsette 1992), as previously described. Larval rearing pans

were placed individually inside Bioquip rearing cages. Adults were permitted to emerge

from the rearing material, and were provided with adult house fly diet and water as

described previously, in the rearing cages, for up to 6 d. After 6 d, all emerged adult flies

were transferred to a transport cage as described below.

In preparation for transport to the field, adult IR house flies were transferred from

the rearing cages to smaller transport cages using CO2 introduced to a cage placed in a

plastic bag. Flies were examined after each application of CO2 for recovery.

Anaesthetized adult flies were gently shaken out of the rearing cage and into a transport

cage. Flies were provided water and adult house fly diet during transport back to the

dairy. The IR adult flies thus produced from dairy-collected-caught larvae ranged in age

from 1-6 d (some adults emerged within hours of collection), depending on the day of

emergence during the week spent in the rearing cage.

After adult flies were removed from the rearing cages and transferred into the

transport cages, the adult house fly diet, water, and larval rearing pans containing dairy-

collected larvae and pupae were placed back into the rearing cages to allow the









emergence of additional IR adult house flies. These rearing pans continued to produce

adult house flies for up to 3 wk.

To further supplement numbers of adult flies, gravid dairy-collected females were

used to produce an Fl generation of adults that were subsequently marked and released at

the dairy. Parental adult house flies were collected in the field by sweep-netting, and

transported back to the USDA fly-rearing laboratory, as described previously.

Oviposition chambers were prepared for each rearing cage and consisted of

approximately 400 ml of 1:1 (v:v) water:GHFD (Hogsette 1992) as described above,

placed in a 473-ml clear plastic deli cup (Model #L2516, Newspring Packaging, Kearny,

NJ). Oviposition chambers were placed inside each rearing cage for up to 3 d to provide

oviposition sites for dairy-collected house flies. After 3 d, the moistened GHFD

containing Fl "dairy-collected" house fly immatures was removed from each oviposition

chamber and added to a fresh mixture of moistened house fly diet in a large 58 x 46 x 8

cm rearing tray (Model #400-3N, Del-Tec Panel Controls, Greenville, SC) by spreading

it evenly across the surface of the fresh diet. This facilitated movement of fly larvae into

the fresh material, eliminating the need to separate larvae from spent material. The

rearing trays were then placed inside a dark-colored close-woven cloth bag, which was

twisted shut tightly and secured with a rubber band to prevent egress of adult house flies.

These covered trays containing the Fl larvae were placed in the previously described

rearing room, and were examined daily until pupation occurred.

After pupation occurred, the pupae were separated from the diet via flotation, then

gently air-dried and separated from chaff in a forced-air chamber (Bailey 1970, Hogsette

1992) for up to 2 h. Finally, the clean, laboratory-produced Fl house fly pupae were









permitted to emerge in a transport cage which already contained 1- to 6-d old IR adult

house flies that had emerged from dairy-collected collected larvae. Because oviposition

chambers were left in the rearing cages for up to 3 d, Fl adults emerged over a 3-d

period. Because daily emergence data were not recorded, the percentages of flies for each

daily age are unknown.

On the next scheduled field release day, cages containing all adult IR and Fl

house flies were transported to the release point. The age and sex of flies that were

marked and released were not recorded.

Transport of Adult Flies to the Field

During transport to the field, food and water were provided ad libidum as

described previously. The transport cage containing adult flies was placed within a 41 x

42 x 35 cm cardboard box (transport box) and covered with a loose-fitting lid. This box

provided shade and was positioned in the center of the vehicle to further decrease

sunlight impact upon the house flies.

Marking, Releasing and Recapturing Adult House Flies

Upon arrival at the release site, the transport cage was removed from the box, and

placed in a sheltered, well-ventilated, and shady location. The transport cage was stored

in this protected location for 2-3 h while alsynite traps were placed in the field as

described below and while additional adult house flies were collected by sweep net. After

all alsynite traps had been placed, adult house flies collected by sweep netting were

added to the transport cage and flies were marked with fluorescent dust in the following

manner.

Corona-magenta (CM) and arc-yellow (AY) Day-Glo fluorescent dust (Day-Glo

Color Corp., Cleveland, OH) were applied on alternating weeks to the caged flies using









metal plunger dusters (Hudson Manufacturing Co., Chicago, IL) to gently pump dust

through the screen. Prior to dusting, the cage was placed inside a large 170-L plastic bag

to simultaneously enhance the application of dust to the flies and reduce the deposition of

dust into the dairy environment. No attempt was made to quantify the amount of dust

applied, or to calculate the amount of dust per fly. Dust was applied until it was visually

apparent that all flies in the cage were well-coated. After the dust was applied, the plastic

bag was tightly closed by twisting and knotting the top. The bagged cage was gently

shaken for 5-10 sec to disperse the dust onto caged house flies. The bagged cage was then

set aside in the shade for up to 2 min to permit the dust to settle inside.

After the dust settled for 2 min, the large bag containing the dusted cage with

marked flies was opened carefully, and downwind of all personnel and vehicles, to

reduce excess transfer of dust to the environment, and to prevent airborne dust from

drifting onto traps without said transfer being performed by active fly transport. The lid

of the cage was lifted slowly and completely to permit egress of dusted adult house flies.

Flies that remained in the cage after 1 min were removed by forcibly tapping the cage

while holding it upside-down directly above the plastic bag. This action resulted in a

secondary dusting for these flies, as they landed in a pile of dust inside the plastic bag.

This pile of dusted flies and excess dust was left undisturbed until all flies had groomed

adequately to permit them to disperse out of the bag and into the dairy environment.

Dispersal of all flies away from the dust pile was completed within 30 min.

Commercially-available alsynite traps (Olson Products Inc., Medina, OH) (Fig. 2-

1) were placed at approximately 0.5-km intervals between the dairy release point and the

adjacent town. Traps consisted of 66 x 33 cm corrugated alsynite panels that were formed









into a 30-cm diameter cylinder. Traps were secured into their cylindrical form with a 2.5-

5.0 cm overlap by insertion of 2-pronged metal fasteners into holes drilled along the edge

of the panels. Traps were inserted into pre-cut slits in either 2 x 2 x 50 cm wooden stakes

(short stakes, provided with trap) or hand-crafted 3 x 3 x 90 cm wooden stakes (long

stakes). Regardless of size, stakes were inserted into the soil until the base of the trap was

approximately 30 cm above ground, to provide a total trap plus stake height of

approximately 65 cm. Sticky Sleeves (Olson Products Inc., Medina, OH), clear plastic

sheets coated with an adhesive and protected with a waxy white peelable paper backing,

were wrapped around the exterior cylindrical portion of the traps, and secured with 2-4

large paper clips. The waxy paper backing was then removed, and the sticky surface was

exposed. Sleeves were labeled with trap number and date of placement on the non-sticky

surface of the clear sheet using a permanent-ink black marker prior to wrapping them

around the alsynite cylinders.

Some traps were placed along the edge of main roads between the dairy and town.

Traps were positioned so that some traps could be used to determine corridor and habitat-

edge movement (Anderson and Danielson 1997, Fried et al. 2005) by placement along

well-travelled roads. Other traps were placed in patchy habitats away from roads. On the

dairies, traps were placed near barns, milking parlors, and calf hutches. In town, traps

were placed close to dumpsters outside two restaurants (traps 12 and 15), two

convenience food stores (traps 17 and 23) and the local post office (trap 16), representing

a non-food site. Four traps (13, 14, 17, and 19) were moved due to difficult access or

human-animal disturbance after wk 2 and renamed (21, 22, 23, 24) so that they were site-

specific. Geographical Information System (GIS) coordinates were recorded at each trap









location and used to generate a diagram of trap placement, and trap locations were

mapped on an aerial image (Google Earth 2009) (Fig. 2-2). Direct distances to each trap

from the release site were determined using the GIS data, and concentric arcs were used

to roughly indicate 0.5 km direct distances. Trapping sites located between the 2.5- and

3.5-km rings from the release site were within the town's border and traps at 3.5 km were

at the far edge of town.

In all instances, attempts were made to ensure that the traps were located as

closely to the 0.5-km concentric arcs as possible. In instances where such placement

involved placing traps on non-public sites, permission was obtained by the respective

homeowner or local business or public school. The purpose of placing traps in multiple

environments allowed for both corridor movement and straight-line flight assessment and

enhanced the probability of recapturing dairy-released, marked house flies. Although it

would be interesting to make the distinction between dispersal due to fly direct flight

versus fly transport on automobiles from the dairy into the town, the data being sought

here were solely to determine if dispersal was occurring rather than to determine how the

dispersal was occurring.

Sticky sleeves were collected weekly, and taken back to the laboratory for

examination under a hand-held 100W long-wave UV light (Model # B-100AP, BLAK-

RAY, Upland, CA). Muscoid flies were identified to genus; house flies and stable flies

were identified to species and the number of fluorescing house flies was recorded.

Dispersal studies were performed from 16 October 2008 to 4 December 2008 and

from 14 May 2009 to 25 June 2009. In 2008, 20 traps were placed in the field weekly for

5 wk, for a total of 5 replications. In 2009, 18 traps were placed in the field weekly for 6









weeks, for a total of 6 replications. Traps were placed in the field on Thursday mornings,

and collected the following Thursday morning. House flies were marked and released

within 1 h after placement of the last trap. Unless otherwise stated, traps were not visited

during the 7 d.

Trap maintenance was conducted so that new sticky sleeves were placed on traps

immediately after the removal of the 7-d exposed sticky sleeves. Collection of used sticky

sleeves was accomplished by placing a protective waxy paper sleeve cover on top of the

adhesive side of the sticky sleeve so that captured flies were protected between sleeve

and cover. This was secured in place with one large paper clip on each end, attaching it to

a cardboard sheet that was cut to the same dimensions as that of the sticky sleeves.

Precautions to minimize secondary transfer of fluorescent dust were taken. These

included covering the sticky sleeves as described. Additionally, researchers involved in

collection of traps used alcohol-based hand-wipes and/or 70% ethyl alcohol, to clean

hands after processing each sticky sleeve. Both of these cleaning methods had proven

successful in removing fluorescent dust from hands and surfaces in preliminary tests

conducted prior to fieldwork. Finally, between field visits, the interior of the vehicle

where traps were stored was wiped down with alcohol-based wipes.

Effects of Fluorescent Dust on House Fly Adults

To evaluate the potentially deleterious impact of fluorescent dust upon dairy-

collected adult house flies, mortality was examined in the laboratory over 24 h under

excessive dusting treatment conditions (Fig. 2-3), hypothesizing that mortality due to

excessive dusting would occur within the first 24 h. Adult house flies were collected

using the aforementioned sweep net at the release site, placed in a transport cage,

provided with food and water ad libidum as described previously, and transported back to









the laboratory. In the laboratory, adult house flies were removed from the transport cage

by anaesthetizing the flies with C02, in a manner similar to that described previously to

transfer flies from rearing to transport cages. The only difference in administration of

CO2 was that the transport cage containing dairy-collected house flies was placed inside a

transport box and lid prior to being enclosed inside the large plastic bag. Because only a

few flies were needed to test mortality impacts of dust treatments, three groups of

approximately 100 anaesthetized flies were removed from the cage by gently scraping a

small index card (62.5 x 75 mm) along the floor of the cage to scoop up the immobile

house flies. Each group was placed in a cardboard ice-cream cup (237 ml, Solo Cup Co.,

Urbana, IL) modified by removing the cardboard base and replacing it with window

screen (940 x 813 microns (16 x 18 mesh); lids were placed upon the ice-cream cup

before flies revived.

The experimental design consisted of three groups of 50 dairy-collected house

flies. Two treatment groups were marked with fluorescent dust (application technique

described below): one group with 0.1 g of corona-magenta dust, and the second group

with 0.1 g of arc-yellow dust. The third group served as a control and was not dusted;

however, the ice cream cup "cage" was shaken in the same manner as if dust had been

added.

Dust was administered by placing the dust on top of the window screen, and

gently pressing it through the window screen with a spoon. A second lid was placed over

the screened base, and the cup plus the two lids were placed inside a self-locking plastic

bag (0.95-L, Ziploc, Racine, Wisconsin), and agitated for 5-10 sec. Dust was allowed to

settle inside the cardboard cups for 5-15 min before opening the bags. Afterwards, the









cups were removed from the bags, and the second lid was removed to reveal the screen

base. Each cup was placed in a plastic deli container with lid, and CO2 was gently

administered without disturbing the dust to achieve knockdown. The flies were then

placed on a sorting tray, where a low-level release of CO2 kept the flies from egressing,

but permitted leg-twitching, while flies were counted into test chambers.

Rectangular pieces of window screen (25 x 20 cm) were folded to form

rectangular test chambers (12.5 cm x 20.5 cm). Test chambers were stapled along two

edges, leaving one short edge open. Ten marked house flies were counted into each test

chamber, and a cotton-ball saturated with 5%-sucrose solution was placed inside to

provide food and water to the flies and to prop the sides of the test chamber apart to

permit fly movement within a 3-dimensional space. Test chambers were maintained in a

laboratory at 33-35% RH, 26-28 C. House fly mortality was assessed 24-h after dust

treatment and flies that were ataxic were considered dead.

Weather

Weather conditions that were observed during the time spent in the study area

were recorded on the days that fieldwork was conducted. To obtain more complete

weather data for the entire study period, weather conditions were obtained from Weather

Underground (2009) at a field recording station located 29 km NE of Dairy A.

Downloaded weather data included daily temperature, relative humidity, precipitation,

barometric pressure, wind speed, and wind direction. Daily data were used to generate

weekly mean values for each climatic factor for the 7-d test period. The 7-d period for

each week began on Friday, and concluded on the following Thursday when traps were

collected, so that weekly mean values were for the 6 d prior to and including the day of

trap collection.









In association with extreme weather conditions, such as approaching storm fronts,

adult fly behavior anomalies were observed. These behavioral anomalies were recorded,

particularly if they appeared to impact house fly dispersal. Additionally, any observations

of unusual larval conditions, such as high mortality due to flooded larval breeding sites

following heavy rains, were also recorded.

Statistical Analysis

Mean, minimum and maximum dispersal distances were calculated using

Microsoft Excel (Excel 2002). Released and recaptured fly data were subjected to PROC

UNIVARIATE to examine normality and PROC MEANS to calculate means using SAS

Version 9.1 (SAS 2002). Paired correlations between numbers of released and recaptured

flies were analyzed using PROC CORR (Pearson's coefficient) (SAS 2002) for both years

combined, for 2008, and for 2009. These data were then submitted to linear regression

analysis with recaptured numbered of house flies regressed against released numbers of

house flies using PROC REG (SAS 2002). These data were regressed for both years

combined, and for each year individually.

Results

The mean dispersal distance for marked flies each week was 0.22 km (range 0.00

- 0.35 km) in 2008 and 0.62 km (range 0.03 1.12 km) in 2009 (Table 2-1). The maximum

weekly dispersal distance ranged from 0.00-1.00 km in 2008 and from 0.10-3.00 km in

2009. Two marked house flies were recovered in the nearby town in 2009 (traps 11 and

15) (Table 2-2). The estimated number of flies that were marked and released each week

ranged from 500-3,700 in 2008 and from 2,000-10,000 in 2009 (Table 2-3). No flies were

marked and released on 7 November 2008 due to a lack of SN-collected flies. No flies

were marked and released on 21 May 2009 due to severe thunderstorms and heavy rain









that prevented collection of house flies. Approximately 9,200 and 28,000 house flies

were marked and released during the 5-wk 2008 test and the 6-wk 2009 test, respectively

(Table 2-3). In 2008, 20 traps collected a total of 13,141 marked and unmarked house

flies; of those, 106 were marked (Table 2-3). The 106 marked flies represented 1.15% of

the 9,200 marked flies released during the 2008 test period (Table 2-3). In 2009, 18 traps

collected a total of 48,435 marked and unmarked house flies; of those, 144 were marked

(Table 2-3). The 144 marked flies represented 0.51% of the 28,000 marked flies released

during the 2009 test period (Table 2-3). Weekly recapture percentages for marked flies

ranged from 0.00 to 1.93% in the 2008 test period and from 0.12 to 0.81% during the

2009 test period (Table 2-3).

Weekly numbers of marked flies that were released varied between 0 in weeks 4

and 7 and 10,000 in week 11. The lowest recapture rate of 0.00% in 2008 followed three

weeks of cold air temperatures that ranged from a weekly average of 7.4 C to a weekly

average of 12.3 C. The preceding two capture weeks also had lower recapture rates as

compared to the recapture following the first two releases. Notably, flies recaptured at the

end of week 4 were part of a release of 500 flies in week 3, so that the 0.20% recapture

rate of week 4 reflects flies that were in the environment in 2008 for 7-14 d (Table 2-3).

The lowest recapture rate of 0.12% in 2009 was obtained at the end of week 7 when no

marked flies were released. These marked flies that were captured on 28 May 2009 were

actually part of a 6,000-fly release made at the beginning of the previous week (week 6)

and had remained in the local environment for 7-14 d (Table 2-3). The highest weekly

recapture rate of 1.93% was obtained in week 2, following release of 3,000 marked flies.









When the cumulative recovery of marked flies over distance from the release site

was calculated, 99.1% (105/106) of the marked flies that had been recaptured were

collected from traps within 0.5 km (<0.5 mi) of the release site in 2008. In 2009, 88.9%

(126/144) and 93.8% (128/144) of the marked flies that had been recaptured were

collected from traps within 0.5 km (<0.5 mi) and 1.5 km (<1.0 mi) of the release site,

respectively (Table 2-2). Total, daily mean and trap mean numbers of all captured house

flies including recaptured marked house flies are reported for each trap in Table 2-4.

Weekly mean weather parameters are shown in Table 2-4. There did not appear to

be any relationship between house fly recapture rates and the weekly mean temperature,

the weekly mean precipitation, or the weekly mean barometric pressure. Dust treatments

did not appear to impact dispersal, as mortality following exposure to AY and CM dust

was 4%, while 2% of control flies died over the 24-h evaluation period.

Production of adult house flies was increased by allowing dairy-collected adult

house flies to oviposit in the house fly diet, resulting in production of approximately

7,000 Fl adults in 2009. Of these, 4,000 were marked and released in wk 3 and 3,000 in

wk 4, representing 40% and 50% of these releases, respectively. Fl progeny adults were

similarly aged (within 3 d) house flies. No Fl progeny were produced in 2008.

The overall number of flies that were released was not correlated to the number of

marked flies recaptured (r=0.72099, p=0.0123). When tested by year, there was still no

correlation between release and recapture rates for either 2008 or 2009. Linear regression

analysis results also indicated no correlation in 2008. However, a positive correlation

existed in 2009 (df=1,4; F=9.28, P=0.0381). Multiple regression analysis for both years

combined showed an overall positive correlation (df=1,9; F=9.74, P=0.0123). The linear









regression line for both years combined is: recapturedflies = 0.402 +

0.0066*released flies.

Discussion

Insects disperse for many reasons, including: to avoid overcrowding, to take

advantage of extended resources available in nearby locations and, to avoid unfavorable

conditions (Stein 1986). Dispersal of house flies and other insects of medical and

veterinary importance needs to be more fully understood, because more than 50% of

infectious diseases world-wide are transmitted by insects (Stein 1986).

Dispersal of house flies, and the impact of fly dispersal on transmission of human

diseases such as epidemic diarrhea, has been scientifically observed for more than 100

years (Kumar and Carmichael 1998). Although the earliest studies are more anecdotal

than quantitative, they nevertheless provide useful clues that have been confirmed

repeatedly using quantitative techniques in the intervening century, regarding house fly

behavior and environmental impacts upon dispersal. For example, Nash (1913) concluded

after his 1904-1909 studies that house flies tend to remain within 0.8 km of their breeding

site in more urban areas, but will readily travel 1.6 km in more rural locations with fewer

human habitations. The tendency of house flies to typically disperse within only 0.8-1.6

km from a breeding or release site has been repeated consistently by multiple researchers

working independently in different geographic locations, unique environmental

conditions, and with different experimental designs (Quarterman et al. 1954, Lysyk and

Axtell 1986, Schoof and Siverly 1954b, Nazni et al. 2005, Winpisinger et al. 2005). This

is consistent with the current study, where the typical distance in 2008 was 0.5 km, and

1.5 km in 2009. For both years combined, >95% of flies were captured on traps placed

less than 1.5 km from the farm.









House flies can disperse over long distances in a short period of time. Pepper

(1944) collected house flies that travelled 5 km in 1.5 d. Bishopp and Laake (1921)

observed that house flies travelled 8.3-10 km, and as far as 21.3 km in 24 h. Shura-Bura

et al. (1956) reported dispersal of 3-4 km within one hour. Sacca (1964) reported house

fly dispersal up to 8 km, while exceptional dispersal distances of >32 km and 33 km were

reported by Schoof and Siverly (1954b) and Murvosh and Thaggard (1966), respectively.

One of the three objectives of this study was to determine if house flies could be

dispersing from the dairy to the town, a 3-km straight-line flight. Because one marked fly

was recovered from a trap in the town, this hypothesis was confirmed by the data. In

general, this short distance is typical for house flies, whose flight range has repeatedly

been reported to be within 1.6 km, despite the exceptional distances travelled by

individual house flies (Schoof and Siverly 1954a,b). Dispersal studies with house flies to

determine the flight range have always been difficult to conduct successfully, especially

in urban settings (Murvosh and Thaggard 1966). Urban communities offer many

substrates that are attractive to house flies, including garbage, decaying grass, coffee

grounds, and excrement from mammals and birds (Schoof et al. 1954b). Schoof and

Siverly (1954b) estimated that an individual house fly may travel 9 km in its lifetime, due

to meandering movements between sites. In my study, traps were placed near garbage

dumpsters outside two restaurants and two convenience stores in town, but not near

residential garbage cans. If residential garbage is attractive to dispersing house flies in the

current study area, then the dispersal data reported in my study would be under-reported.

In my study, dispersal of house flies was observed up to a maximum direct-line

distance of 3 km over a 7-d test period. However, because I collected traps only weekly, I









was unable to determine dispersal rate. It would be informative to repeat this study with

collection of sticky traps over shorter intervals, such as daily or hourly, to obtain detailed

information about the temporal dispersal rates of house flies from dairies into this north-

central Florida town. It would also be useful to track fly meandering movement between

sites by placing self-marking traps (Hogsette 1983) with a unique color of dust at each

site. By doing so, recaptured flies that possess more than one color of dust would clearly

have been present at multiple sites between release and recapture.

Although house flies may disperse away from a breeding site, their dispersal may

be more complicated than merely traveling from point A to point B. There is evidence to

substantiate observations that flies disperse with "randomized, reciprocal type

meanderings" (Williams 1973) that result in back and forth travel between the

breeding/release site and some other site(s) of interest as distant as 17 km (Schoof and

Siverly 1954b). This type of dispersal is of particular importance in areas where large

animal-rearing facilities such as dairies with endemic pathogenic E. coli 0157:H7 are

located within close proximity to human populations because it could greatly increase the

potential for pathogen transmission to humans. Because flies were only marked at the

release site, I was unable to determine if captured house flies had been at an alternative

attractant site other than the dairy release site. Future studies could examine the

meandering behavior of house flies in this study area by release of differently colored

marked flies at multiple locations within the study area. For example, the dusting of flies

at several dairies and at the garbage dumpsters in town would allow observation of

dispersal patterns within the entire study area.









Dispersal of house flies may be complicated by behavioral tendencies of insects to

use landscape features to facilitate movement along corridors or to follow habitat-edge

environments (Anderson and Danielson 1997, Fried et al. 2005, Reisen 2010).

Furthermore, distances travelled by dispersing house flies are typically measured as direct

flights from point A to point B, with no regard to potential fly dispersal along corridors

such as highways and/or vehicle-assisted movement (Nazni et al. 2005). If flies use

highways as corridors for undirected dispersal movement, then distances travelled could

actually be under-representative of the capability of flies that travel directly to new sites.

Interestingly, Quarterman et al. (1954) report that house flies have a "living space" of up

to 6 km diam over which they roam freely. Flies travel randomly throughout this space

and aggregate at multiple feeding and breeding sites so the populations at multiple sites

becomes one huge metapopulation. Quarterman et al. (1954) also observed that house

flies disperse rapidly away from the release site, and that trap catches were highest on the

first 3 d following releases. Similarly, Schoof and Siverly (1954b) demonstrated that

house flies display random dispersal and directional reversals by using three secondary

mark and release sites in addition to the primary release site. Their release of radio-

labeled flies from a primary site followed by dusting and release and recapture of these

radio-labeled flies at three secondary sites indicated movement in all directions between

the multiple sites.

Interestingly, my study indicates that house flies dispersed by both direct overland

flight and by corridor movement. Direct flight is strongly implicated by the recapture of

many flies at a neighboring dairy located 1.5 km to the west, with no direct road access

connecting the two dairies. These two dairies were separated only by a common fence









line and patchy habitat that included tree wind breaks between their mostly open pastures

that held cattle. Similar dispersal of house flies from one dairy to an adjacent dairy was

also observed by Denholm et al. (1985) and Lysyk et al. (1986).

House fly movement along landscape corridors with barriers, such as tree lines, is

also likely (Fried et al. 2005), and can occur by both vehicle-assistance and by direct

flight. Vehicle-assistance is likely due to milk and feed trucks, and other service vehicles

that visited both dairies on multiple occasions during the field study. Similarly, Sacca

(1964), Nazni et al. (2005) and Sievart et al. (2006) reported passive transportation of

flies between dairies by automobiles. House flies were frequently observed on the

exterior of the research vehicle after departing the release site, for up to 5 min. To reduce

the potential for providing vehicle-assisted transport to the flies, typical departure from

the release site was followed by driving away from the town along roads for up to 15 min

until no flies were observed on the vehicle. During this time, efforts were taken to rid the

interior of the vehicle of any flies. Additionally, there was no travel in this study towards

the town after releasing the marked flies. It is unknown what impact the milk and feed

trucks, and other vehicles had on fly dispersal between the release site and the town.

Using only traps located 0.5 3.5 km, and excluding the four traps that were

removed from the study after wk-2, the number and percentage of traps that were positive

for recovery of a marked house fly were approximately equal for both roadside (5/8 traps,

62.5%) and patchy-habitat (7/11 traps, 63.6%) traps. Although the number of positive

traps was very similar, the roadside traps recovered higher numbers (n=39) of marked

flies than patch-habitat traps (n=16). This suggests that flight by house flies along

corridors (Fried et al. 2005, Reisen 2010), as reported for other insects, is more important









than flight through patchy-habitats in this region of north-central Florida. However, it

also clearly indicates that both types of dispersal occurred, and that marked flies were

disseminated throughout and active at multiple sites throughout the study area. Further

dispersal by recaptured individuals might have occurred had they not been prevented

from continued travel by the adhesive sticky sleeve.

One marked house fly was recovered at a trap placed at Restaurant C, located at

the maximum recapture distance (3.0 km) obtained in this study. Although Restaurant C

was located in a tree-covered patchy habitat, it was also located close to the major road

that ran east-west through town. Therefore, no conclusions can be made regarding the

probable dispersal method or direction of approach used by the fly that was recovered at

Restaurant C (trap 15). The fly could have arrived there by undirected flight, visiting

multiple attractive sites along the way (Schoof and Siverly 1954b), by following

corridors provided by roads (Johnson 1966), or by direct flight through-the-woods flight

(MacLeod and Donnelly 1960).

Marked flies were recaptured every week except week 5 (Table 2-3), which

followed several weeks of lower temperatures (7.2 to 9.5 OC) than surrounding weeks

(Table 2-4). Dispersal in the study area occurred consistently with average weekly

temperatures that ranged from 12.9 to 25.7 C. This was not unexpected because the

house fly flight temperature threshold is approximately 13 C (Taylor 1963). Although

the last three weeks in 2008 had weekly average temperatures above the house fly

activity threshold, it is possible that the accumulation of low temperatures and shorter

daylight hours inhibited dispersal behavior. This is reflected in the very low recapture









rates following week 3 and 5 releases. Because week 5 was 21 d in duration, the last three

collections of the 2008 study period represented a 5-wk calendar period.

A further challenge of dispersal studies is the diffusion (dilution) of marked

insects across a landscape as the distance from the release site increases (Schoof and

Siverly 1954b, Stein 1986). The capture of one marked fly 3 km from the release site

could be equivalent to capture of several house flies at the release site, because the

density of traps present at more distant concentric distances was less over a greater

geographic area per trap than traps located more closely to the release site. Schurrer et al.

(2004) determined that captures of house flies diminished with increased distance from

the release site. House flies are attracted to a wide variety of substrates, and every time a

fly lands on an attractive substrate, the number of flies moving away from the release site

is diminished (Murvosh and Thaggard 1966). Schoof and Siverly (1954b) estimated that

approximately 50% of flies that land on an attractive site might subsequently progress

towards an urban community that contains numerous attractants, so that the number of

flies decreases by half at each visited site between the dairy and the town.

The fact that such a small number of flies were recovered in town is probably a

result of the manner in which the numbers of marked flies spread out spatially, thereby

diluting the likelihood of recapture as they moved further from the release site

(Quarterman et al. 1954, Schurrer et al. 2004, Krafsur et al. 2005). An estimated total of

37,200 marked flies were released during this study. The release of more marked flies, in

theory, should increase the likelihood of capturing a marked fly on the far side of the

town. Thimijan et al. (1972) increased their recapture rates from 0.17% (3 traps) to 0.51%

(44 traps); however, their study was conducted in a closed barn so that an increased









number of traps would likely influence recapture rates in their study more than placement

of additional traps in my 3 km outdoor study area.

If the marked flies in this study were dispersing from the release site in all

directions as reported in previous studies (MacLeod and Donnelly 1960, Pickens et al.

1967), then the numbers of flies moving towards the town after a release represents only

a portion of the released flies. I placed traps and recaptured marked flies in only one

eighth (SSW portion) of a full 3600 circle, to study the dispersal of flies from the dairy

into town. If the flies were dispersing equally in all directions (undirected), then 106 and

144 recaptured flies in 2008 and 2009 respectively, would be equivalent to a recapture

rate of 848 (9.2%) and 1,152 (4.1%) in all eight ordinal directions. Dispersal of the house

flies in multiple directions was very likely in the current study area, because there are

several nearby dairies and poultry farms surrounding the release dairy. If dispersal of flies

was nondirectional, then attractive sites in town that are sufficiently attractive to interrupt

flight might receive a disproportionate number of flies from the dairies (MacLeod and

Donnelly 1960.

The effect of wind speed and direction upon house fly dispersal has been

examined with varying results by previous researchers. Flies have been reported to be

blown long distances. Bishopp and Laake (1921) cite wind-assisted dispersal of flies for

distances ranging from 3.6 km (Hodge 1913) to a remarkable distance of 153 km (Ball

1917). During this study, monthly prevailing wind directions were southerly overall for

the entire study. In 2008, prevailing winds were initially SSE and changed to SSW. In

2009, prevailing winds for June, July, August, and September were W, SW, SSW, and

SE, respectively (Weather Underground 2009). Although in coastal areas of Florida, sea-









breeze fronts can influence wind direction, central Florida is too far from coastal fronts to

have any influence most of the time (Jones et al. 1991). The town was SW of the dairy

release site, so for all months, fly dispersal was at least partially into the wind. In this

situation, fly dispersal from the dairy into town would not passively rely on wind.

Evidence that flies actively travel into the wind or at right angles to the wind

using olfactory and optomotor senses for direct flight, rather than being passively or

actively transported by the wind was noted by Bishopp and Laake (1921) and Johnson

(1966). Such movement would have resulted in flies dispersing towards the town.

The discrepancy between directed and undirected dispersal concepts suggests that a

better-designed study on the potential of fly pathogen transmission into communities is

needed to implement improved fly control programs at animal-rearing facilities. The

scope of such a study would need to include all house fly-producing facilities within 6

km. However, each of those facilities also would share a fly population with additional

sites located within another 6 km distance, making such a study impractical and perhaps

impossible.

Fly density could have been a dispersal factor in my study, because increased

population density leads to increased displacement (Stein 1986), and by laboratory-

rearing of dairy-collected progeny followed by release of those flies at the dairy, the

density of the house fly population may have been artificially increased. However, adult

fly density on the dairy was not estimated, so the effect of population density upon

recapture rates cannot be determined in the current study. When recaptured numbers of

flies were regressed against the number of released flies, there was a linear relationship

for both years combined, and for 2009. However, there was no relationship in 2008. This









implies that the numbers of flies that were released, including the laboratory-reared flies,

might have influenced the overall dispersal rates or patterns of the representative fly

population in 2009, but not in 2008.

Capture of marked house flies 8- to 14-d after traps were placed demonstrates the

ability of some marked flies to survive for longer than 7 d (Table 2-3), which confirms

the findings of previous studies (Lindquist et al. 1951; Schoof and Siverly, 1954a,b;

Pickens et al. 1967). Recapture of marked flies for up to 8-14 d after their release also

provides information for how long any adult fly, whether marked or not, lives in the

environment. This is probably highly variable and is not known with certainty. Survival

of marked flies for more than 7 d also implies that flies might disperse from breeding and

aggregation sites on different days during their lifetime. While placement of traps for a 7

d period might be sufficient to capture most marked flies, it clearly is not long enough in

all cases, as adult-marked flies have been recaptured 10 d (Schoof and Siverly 1954a) and

20 d (Lindquist et al. 1951) after release, and pupal-emergent marked flies have been

recaptured 18 d after release (Pickens et al. 1967). In the current study, survival of

marked house flies for more than 7 d was only recorded during weeks 4 and 7, so most

house fly dispersal likely occurred within 7 d after release. This is consistent with 7-d

mean recovery rates of 84.7% and 95.5% in two field trials conducted by Pickens et al.

(1967). Because most house flies seem to disperse within 7 d, it is likely that the principle

potential health threat posed by house fly transmission of disease-causing pathogens

might be heavily concentrated within this time period.

However, the ability for a few flies to survive for longer periods, combined with

their continuous, random dispersal (Schoof 1959) from a release site to adjacent dairies









and into urban areas could result in successful pathogen transmission over an extended

period of time, so that control of house fly populations should include measures that are

effective for longer than 7 d. In my study, flies that had been in the environment for

longer than 7 d were recaptured in wk 4 (7 November 2008) and wk 7 (21 May 2009).

Therefore, survival in the environment past 7 d does not appear to be limited by season

(Table 2-1). Daily replacement of trap sleeves over a 2-wk period would provide more

thorough information for daily dispersal rates and total recapture percentages. Increased

trapping periods should be accompanied by incorporation of additional dust colors, so

any flies that might survive 15 d or more could be observed.

Flies should be caught, marked and released early enough in the day to promote

fly activity, including dispersal. Pickens et al. (1967) observed that marked house flies

tended to rest for 15-30 min on buildings, fence posts and trees near the release site

before dispersing. If releases are done too late in the day, then flies may not groom

adequately or disperse due to decreased metabolism and avoidance of nighttime flight.

Flies in this study were released at approximately 3:00 p.m.

The weekly number of marked and released house flies ranged from 0 (no release

due to low fly numbers or weather conditions) to 10,000. The total number of flies

marked and released in this study (37,200) was low in comparison to those of previous

mark-release-recapture studies: e.g., 171,427 (Schoof and Siverly 1954b) and 160,000

(Nazni et al. 2005). However, my weekly release levels were similar to Quarterman et al.

(1954) who marked and released 13,500 flies, and Thimijan et al. (1972) who marked and

released 5 groups of 2,500-5000 flies. However, the test area used by Thimijan et al.

(1972) was a closed barn (17 x 10 x 3 m) that restricted fly dispersal. This study's 0.50%









marked house fly recapture rates are higher than the filth fly recapture rates of 0.014-

0.029% (mean 0.022%) obtained by Tsuda et al. (2009).

When examining the Yates et al. (1952) data following exclusion of their highest-

capture trap which was placed adjacent to the release site, it is interesting to note that I

obtained higher per trap recapture rates with placement of traps in only one ordinal

direction (SW) away from the release site. Yates et al. (1952) placed their traps in eight

ordinal directions. One can speculate from the relatively high mean recapture rates per

trap during my week of highest release (wk 9) that house flies dispersed directionally

towards the town. The presence of SSW prevailing winds during wk 9 further strengthens

this hypothesis, because flies that dispersed towards town during wk 9 dispersed into the

wind. This in turn implies that the role of house flies in potential pathogen transmission

from dairies into urban communities might be more important than currently recognized.

Overall prevailing winds during the entire study were southerly, so that dispersal of

recaptured flies was probably not wind-assisted, but attractant-seeking or odor-driven

instead. Identification by dispersing flies of attractive sites in urban communities is likely

a factor in house fly dispersal into these areas.

Dispersal distances and recapture rates might be influenced by the type of flies

used, i.e., field-collected or laboratory-reared, as mentioned briefly above. Previous

studies have indicated that use of field-collected flies is more representative of dispersal

under natural conditions than flies that are reared for multiple-generations in the

laboratory. Eddy et al. (1962) recaptured a 10-fold higher percentage of field-collected

flies than laboratory-reared flies, implying that laboratory colonies may lose the ability to

disperse. Although laboratory-reared flies might lose the ability to disperse, their









introduction in large numbers might increase the population of adult flies, causing

increased dispersal of flies from the dairy, either of the natural population or of the newly

introduced flies. It would be interesting to study the effect of large releases of flies upon

existing house fly populations, to determine if natural dispersal rates match dispersal

rates of recaptured marked flies.

There were weather conditions that made collection of adult flies very easy, such

as on days that storms were approaching when adult flies were very active in the air and

remained hovering in the air in large numbers, even without human disturbance. This

corresponds with previously reported accounts that fly activity greatly increases under

falling barometric pressure conditions associated with storm fronts (Wellington 1945,

Holzapfel and Harrell 1968). Although there were no appreciable changes in overall

barometric pressure (BP) during the entire study period, it is possible that changes in the

barometric pressure in the hours preceding release of marked flies might have influenced

house fly dispersal in this study. Barometric pressure decreases in an undulating fashion

and can only be observed with a column of mercury (Wellington 1945); thus, a net

change in barometric pressure might be slight, yet still influence fly behavior. However,

hourly weather data were not available, so that examination of the effect of weather for

periods of less than 24 h was not possible.

On 21 May 2009, the study area was experiencing its fifth consecutive day of

steady, heavy rain that resulted in extensive flooding. Adult fly activity was negligible on

this date. On 4 June 2009, fly behavior at both dairies consisted of hovering in large

clouds approximately 2-3 m above the ground instead of the usual feeding and resting

behavior. The daily average weather data obtained from the nearest weather station does









not support these field observations. This is very likely due to the fact that the weather

station was located approximately 29 km inland. Proper determination of weather

influences upon house fly behavior should be tracked locally in future studies, as distant

weather stations do not provide data that always accurately responds to local

microclimates.

In Florida, adult house flies are primarily active mid-day during spring and fall

when maximum temperatures are cooler (Hogsette, personal communication). They adopt

a diurnal activity behavior with increased summer temperatures, so that they become

inactive during the hottest part of the day, and have two activity peaks: the first peak

occurs in the morning and the second peak before dusk. During the 2009 portion of this

study period, the daily temperatures steadily increased necessitating an earlier sweep-net

collection before the temperatures became hot enough to inhibit fly activity. Because the

hot weather decreased adult house fly activity, another obstacle for sweep netting

collection was created by flies congregating near the feed troughs that lined the edges of

the barn. As the temperature became progressively hotter, the cattle spent increasingly

longer periods in the shaded protection of the barn that also was equipped with large

ceiling fans and mist sprayers that cooled the animals. When cattle were present in the

barn, they had ad libitum access to the feed troughs, and therefore frequently had their

heads in the grain where flies were located, making sweep netting difficult. Because

cattle are easily frightened by sudden movements, in the interests of cattle safety, flies

were not sweep netted at the feed troughs when cattle were present.

The collection and rearing of immature flies and F rearing of adult flies greatly

increased the number of house flies available for mark and release. In particular,









laboratory-rearing enabled the release of more uniform numbers of flies even when field

populations were at relatively low levels. Release of mass numbers of laboratory-

produced F dairy-collected adult flies could have changed dispersal behaviors for a

couple of reasons. Increased larval density in the laboratory could produce smaller adults

(Haupt and Busvine 1968), increase developmental time (Black and Krafsur 1986), or

decrease adult emergence. These factors could delay adult emergence of earlier instars so

that multiple age-groups of flies entered the adult population simultaneously. Previous

authors have noted that overcrowding led to decreased food consumption resulting in

smaller adult sizes (Haupt and Busvine 1968) with increased activity, and increased

dispersal tendencies (Johnson 1969, Stein 1986). Smaller adult size may influence sexual

maturity as well, so that small individuals might not be sexually mature. Because sexual

maturation of the ovaries inhibits or decreases the dispersal tendencies of the adult female

fly (Johnson 1969, Stein 1986), it is possible that small stature could prolong the

dispersal period past the typical 3 d previously reported for the house fly. Similar

observations of reduced size and increased activity were previously described by Taylor

and Sokal (1976), who also stated that larval overcrowding increases the tendency of

adults to disperse.

Initial efforts to maximize production of adult house flies from dairy substrates

containing immature fly stages of all ages were very time-consuming and inefficient,

because of the emergence of adults in small numbers over a 1-3 wk time period.

Allowing dairy-collected adult house flies to oviposit in the house fly diet increased the

production of same-age house flies that could be mass-reared using well-established

standard USDA house fly rearing procedures (Hogsette 1992).









Provided that laboratory-rearing did not alter oviposition behavior or larval

mortality, the F generation produced by dairy-collected adult flies should accurately

represent the real fly population's relative mixture of sexes. However, age might not be

accurately represented because natural populations have uneven distribution of different

larval stages in the spilled grains at the dairy (Johnson 1966), whereas oviposition of

dairy-collected females generated a same-age population that was 1-3 d post-eclosion.

During the rearing process, larvae were observed clumping together in groups comprised

largely of same-age instars. This behavior could indicate different nutritional or

environmental needs or changing feeding capabilities of each life stage. However, this

observed behavior in the laboratory might be changed from their behavior in the field,

because they were reared under a different light:dark regime and were fed a different diet.

One example of changing environmental needs for developing house fly larvae is seen

when third instars migrate before pupating. While feeding larvae are active throughout

relatively moist grain, aggregates of pupae are found in drier sections of the grain. Dairy-

collected house flies were observed to pupate just under the crust of spilled grains at the

dairy. In the USDA laboratory-rearing facility, IR and Fl larvae mostly migrated

completely out of the tray, and pupated in the folds of the cotton pillowcase used to cover

the trays. Many larvae were also clumped together in the covers of the rearing trays.

My dispersal study also may have been influenced by the age of released flies.

Johnson (1966, 1969) observed that younger flies disperse more readily than older flies.

Because laboratory-reared progeny of dairy-collected flies were young flies aged 1-6 d,

their dispersal activity might be higher than that of the typical dairy farm flies in the

study area. In apparent contrast to Johnson (1966), Taylor and Sokal (1976) observed that









older flies (3-6 d) dispersed readily while younger flies (1-3 d) did not disperse.

However, Johnson (1966) noted that muscid flies disperse inter-reproductively, i.e., after

ovaries have matured, as well as pre-reproductively, and that females are more likely to

disperse before egg development occurs, while gravid females are more likely to cease

flight, and to land on suitable oviposition sites. Similarly, Sasaki et al. (2000) observed

that female flies aged 6-8 d were more dispersive than 6-8 d old males. On a different

note, Johnson (1966) states that populations, rather than dispersing, tend to mix within

wide areas. Evidence of this is seen in the current study, because marked flies were

recaptured at neighboring farms.

On the one hand, the laboratory rearing methods used in this study may have

produced well-fed adults, as the nutrition level could have been greater in the GHFD

media than in the dairy's spilled grains. This might have resulted in "fatter" or healthier

flies that may not have been inclined to disperse. Larval weights were not measured, so it

cannot be determined which situation might have prevailed for either laboratory-reared or

dairy-collected flies. Furthermore, it is possible that conditions changed from week to

week in the laboratory-reared flies, because the density of fly larvae in the GHFD was not

measured. Eddy (1962) observed that laboratory-reared flies do not disperse as readily as

field-collected flies. However, Pickens et al. (1967) did not observe any difference in

dispersal rates or patterns between field-collected and laboratory-reared flies.

Conversely, larval crowding may have occurred in the trays, so that the house fly

larvae were underfed and subsequently produced undersized adult flies (Black and

Krafsur 1986). Although I did not record fly weights or sizes for any fly stages used in

this study, I did observe larval overcrowding during production ofF1 field-collected









progeny for release on 4 June 2009. During week 2, the F progeny exhausted the GHFD

media, and half of each tray's larvae and exhausted media were transferred into clean

trays containing fresh GHFD. Exhausted media containing flies was spread on top of

fresh GHFD. The flies moved into the new media. The adult flies resulting from this

production took longer to develop (19 d) than the USDA colony flies (14 d), appeared

smaller, and were noticeably more active than same-age laboratory colony flies (personal

observation), which were maintained in an adjacent USDA rearing cage. Therefore, the

laboratory-rearing methods used in this study probably served, in at least one release

week, to increase the dispersal behavior of the IR and Fl flies. This could have occurred

if population pressure was placed on the house fly populations upon release due to mass-

introduction of thousands of flies. It might also have occurred because the individual flies

were more inclined to disperse due to effects of larval overcrowding (Taylor and Sokal

1976, Black and Krafsur 1986). Increased dispersal rates due to these factors may have

contributed to the high 0.81% (81/10,000) recapture rate of flies that had been released on

4 June 2009, when 40% of the released flies were Fl laboratory-reared progeny.

Conversely, transport of laboratory-reared progeny to the field in cages containing mixed

sexes of flies aged 1-6 d with food and water provided ad libitum may have decreased

dispersal tendencies, as many flies may have mated and obtained protein meals needed

for egg development.

Environmental conditions such as temperature, precipitation, barometric pressure,

wind speed and wind direction were evaluated individually to determine their impact on

the dispersal of marked house flies. There may be a relationship between the number of

flies that were marked and released and the recapture rate, although these data were not









statistically analyzed in this study. There does not appear to be a relationship between

precipitation and recapture rates. These results agree with those of previous researchers

who also did not find a correlation between weather and house fly behavior (Eddy 1962,

Lysyk 1993).

Under natural conditions, there are many age groups of adult house flies at

breeding and aggregation sites. It is unlikely that all adult flies disperse from these sites,

and dispersal may be dependent upon house fly sex and age. The female to male ratio of

adult flies collected in sweep nets may have influenced the dispersal rates and patterns,

and female and male adults may rest in different preferential sites during the day

(Avancini and Silveira 2000); therefore, sweep netting in mid-day over feed troughs may

have resulted in predominant collection of females.

Events occurred on the dairy that influenced fly populations sizes, larval

development, and behavior. Mechanical fly control efforts included sanitation and habitat

elimination, such as the removal of spilled grains, drainage of manure lagoons, and

application of manure slurry to crop-fields. Chemical control efforts included frequent

application of permethrin to the backs of cattle, intermittent application of imidacloprid-

containing fly baits and intermittent application of permethrin around the exterior of the

barns and milking parlors. The dairy also utilized pyrethroid-impregnated ear tags on the

animals. However, ear tags are unlikely to have decreased house fly abundance on the

dairy due to fly biology and behavior.

Several interesting results were observed in this study: 1) house flies dispersed 3

km from a dairy to a restaurant in a nearby town; 2) recapture rates (1.15% in 2008 and

0.50% in 2009) were comparatively high in this study; 3) recapture occurred at sites









located within patchy habitats, implying direct flight over obstacles or wind-assisted

dispersal; 4) recapture occurred at sites along major roads that provide corridor

movement, implying possible landmark orientation or vehicle-assisted transport; 5) most

marked flies were recaptured within 7 d; and, 6) recapture of one fly (0.2%) in week 4

and seven flies (0.12%) in week 7, despite zero-releases the previous week, document fly

survival and dust retention for between 8 and 14 d.

The numbers of house flies recaptured in this study are probably under-reported

because house flies disperse in all directions (Pickens et al. 1967). The potential for house

fly transmission of pathogens from dairies into town is clearly demonstrated by recapture

of at least one fly at a restaurant in town. Because dairy cattle are the primary reservoir

for enteric pathogens such as E. coli 0157:H7 (Dunn et al. 2004b, Nmorsi et al. 2007),

and this pathogen can remain viable on house fly exteriors (De Jesus et al. 2004) and in

house fly guts for up to 4 d (Kobayashi et al. 1999, Sasaki et al. 2000), the potential for

disease occurrence is tremendously increased when house flies successfully disperse into

communities from dairies. In that dairies provide both the source and dispersal

mechanism for pathogen transmission, this study shows the importance of pathogen and

house fly management that dairy operators should consider. This also documents the need

for additional efforts to educate and encourage producer utilization of available control

methods to decrease house fly populations.














































Figure 2-1. Alsynite trap (Olson Products Inc., Medina, OH) placed at dairies and used to
recapture on-dairy and dispersing house flies.















122

































Figure 2-2. Alsynite trap (Olson Products Inc., Medina, OH) locations and distance (km) from
release point. Trap 1 is located at the release point. Traps that are located within less
than 0.25 km of a concentric circle are considered to be located at that radial distance.
Trap distances are as follows: Traps 3, 4 and 5: 0.5 km; 6 and 20: 1.0 km; 7, 8, 24,
and 25: 1.5 km; 9 and 21: 1.75 km; 10, 18 and 22: 2.0 km; 11, 17 and 23: 2.5 km; 13,
15 and 16: 3.0 km; 12 and 14: 3.5 km. Traps are placed at approximately 0.5 km
intervals radiating out from the release site at the dairy, to the town which is located
W-SW of the dairy. Slight displacements necessary to accommodate roads, private
property, and other obstacles. Traps placed along major corridors such as roads, in
edge habitats between open fields, and along shrub/tree lines.

























A









',* a .

*... S






** **. .* *







S..... ..
*** B


















Figure 2-3. Examples of dairy-collected-collected house flies following excessive treatment with
two dusts to determine 24 h mortality effects. A) flies dusted with corona-magenta
dust, B) flies dusted with arc-yellow dust, and C) flies not dusted (control).









Table 2-1. Mean and maximum distances flown per week and year by marked house flies
released at a dairy in north central Florida.


Distance (km)
Mean
0.01
0.29
0.35
0.00
NA
0.22
0.39
0.73
1.12
0.73
0.10
0.64
0.62


Maximum
0.10
1.00
0.50
0.00
NA
1.00
1.50
2.50
1.75
3.00
0.10
1.50
3.00


a Flies were collected during two study periods:
and wk 6-11 (21 May 2009 to 25 June 2009).
b Wk 4, marked flies recaptured only at release ,
c Wk 5, no marked flies recaptured (NA, not apj


wk 1-5 (23 October 2008 to 4 December 2008)


Week
1
2
3
4b
5c
2008 Total
6
7
8
9
10
11
2009 Total











Table 2-2. Trap distances (km) from the release site, the total numbers of marked house flies captured per alsynite sticky trap, and the
cumulative percentage of marked flies captured across all study weeks from the dairy release site to the more distant traps
placed in a rural north central Florida landscape.
2008 2009
Test (wk) Test (wk)


Distance Trap
from No.b
Release
Site (km)
0 1
0.1 2
0.5 3


1.0


1.5


1.75

2.0



2.5


3.0


1 2 3
17 1 0
27 56 2
0 0 0
0 0 1
0 0 0
0 0 0
0 0 0
0 1 0
0 0 0
NAd NA 0
NA NA NA
0 0 0
NA NA 0
0 0 0
0 0 0
0 0 NA
NA NA 0
0 0 0
0 0 NA
NA NA 0
0 0 NA
0 0 0


4 5
1 0
0 0
0 0
0 0
0 0
0 0
0 0
0 0
0 0
0 0
NA NA
0 0
0 0
0 0
0 0
NA NA
0 0
0 0
NA NA
0 0
NA NA
0 0


Marked
Flies
19
85
0
1
0
0
0
1
0
0
NA
0
0
0
0
0
0
0
0
0
NA
0


C

Fl
1
9

9


um.
%
iesb 6 7
7.9 11 1
8.1 0 4
- 25 0
9.1 0 0
0 0
0 0
0 0
)0.0 0 0
0 0
1 0
1 1
0 0
0 0
0 0
NA NA
NA NA
0 0
0 1
NA NA
0 0
NA NA
0 0


8 9 10 11
0 52 6 1
1 11 3 2
0 9 0 0
0 1 0 0
0 0 0 1
0 0 0 1
0 0 0 0
0 0 0 0d
0 1 0 0
0 1 0 1
1 0 0 0
0 0 0 0
1 4 0 0
0 1 0 0
NA NA NA NA
NA NA NA NA
Of 0 0 0
0 0 0 0
NA NA NA NA
0 0 0 0
NA NA NA NA
Of 1 0 0


Cum.
%
Fliesb
51.1
63.8
88.9


92.8


93.8


97.2

97.9



100.0


Marked
Flies
71
21
34
1
1
1
0
0
1
3
3
0
5
1
NA
NA
0
1
NA
0
NA
1


1(











2008 2009
Test (wk) Test (wk)
Distance Trap
from No. b Cum. I Cum.
Release Marked % Marked %
Site (km) 1 2 3 4 5 Flies Fliescb 6 7 8 9 10 11 Flies Fliesb
16 0 0 0 0 0 0 NA NA NA NA NA NA NA
3.5 12 0 0 0 0 0 0 0 0 0 0 0 0 0 100.0
14 0 0 NA NA NA NA NA NA NNA NA A NAA NA NA
Total 44 58 3 1 0 106 100.0 38 7 3 81 9 6 144 100.0

aFlies were collected during two periods: wk 1-5 (23 October 2008 to 4 December 2008) and wk 6-11 (21 May 2009 to 25 June 2009).
b Some traps were placed along main roads (3, 6, 7, 9, 10, 11, 22), while others were located in patchy habitats (4, 5, 8, 12, 13, 14, 15,
16, 17, 18, 19, 20, 21, 23, 24, 25). Traps 1 and 2 were located outside opposing ends of release farms feed barn.
' Cumulative percentage of marked flies captured from the release site to more distant traps. Traps (11-17, 23) located at > 2.5 km
were located within the town. Traps not in town were considered rural.
d Trap 20 was missing.
e NA, not applicable. Traps not used for collection were eliminated from any analysis.
f Traps fell onto their sides on the ground.











Table 2-3. Weekly recapture rate of marked house flies on alsynite sticky traps, as a percentage of the number dusted and released the
previous week at a dairy in north central Florida.
Release No. Dusted Dusted Collection No. Marked Recaptu
Test (wk)a Date & Released %SN:IR:F1 b Colorc Date Duration (d) Flies red %
Collected Color
Recapture
1 10/16/08 3,700 100:0:0 AY 10/23/08 7 44 AY 0.08
2 10/23/08 3,000 100:0:0 CM 10/30/08 7 58 CM 1.93
3 11/01/08 500 100:0:0 AY 11/07/08 6 3 AY 0.60
4 11/07/08 0 NA NAd 11/13/08 6f 1 AY 0.20
5 11/13/08 2,000 100:0:0 CM 12/04/08 21 0 CM 0.00
2008 Total 9,200 100:0:0 NA NA 106 NA 1.15
6 05/14/09 6,000 100:0:0 AY 05/21/09 7 38 AY 0.63
7 05/21/09 0 NA NA 05/28/09 7f 7 AY 0.12
8 05/28/09 2,000 50:50:0 CM 06/04/09 7 3 CM 0.15
9 06/04/09 10,000 60:0:40 AY 06/11/09 7 81 AY 0.81
10 06/11/09 6,000 50:0:50 CM 06/18/09 7 9 CM 0.15
11 06/18/09 4,000 100:0:0 AY 06/25/09 7 6 AY 0.15
2009 Total 28,000 71:4:25 NA NA 144 NA 0.51


NA, not applicable.
a Flies were collected as follows: wk


1-5 (23 October 2008 to 4 December 2008) and wk 6-11 (21 May 2009 to 25 June 2009).


b Estimated percentages of flies marked with fluorescent dust and released by each of three methods described in the text. SN, adult
flies were sweep net captured, marked and released same day; IR, Immature flies were dairy-captured and laboratory-reared, then
marked and released after adult emergence; F1, progeny were laboratory-reared from ovipositing dairy-collected adults, then marked
and released after adult emergence.
' Dust color: AY = arc-yellow; CM = corona-magenta; NA = no dust applied.
d Percent recapture = (No. marked house flies recaptured divided by the no. house flies dusted and released) 100.
SNo flies were marked and released in week 4 or in week 7.
f Flies collected in weeks 4 and 7 were recaptured from previous releases in weeks 3 and 6 respectively. These data were not used in
any analysis.











Table 2-4. Total number of house flies and marked house flies captured on alsynite sticky traps following release at a dairy farm in
north central Florida. Capture data are normalized to number of flies per trap per day to adjust for different collection
period lengths.


Wk Date Collected
1 10/23/2008
2 10/30/2008
3 11/7/2008
4 11/13/2008
5 12/4/2008
2008 Total
6 5/21/2009
7 5/28/2009
8 6/4/2009
9 6/11/2009
10 6/18/2009
11 6/25/2009
2009 Total
Grand Total


Durationb (d)
7
7
6
12
21

7
14
7
7
7
7


Total House Flies Captureda
Daily Mean' Trap Meand
Total (flies/day) (flies/trap/day)
4,655 665 33
3,096 442 22
2,847 475 79
1,702 142 14
841 40 2
13,141 353 30
15,946 2,278 127
12,632 902 100
3,101 443 25
9,697 1,385 77
2,867 410 23
4,192 599 33
48,435 1,003 64
61,576


Marked House Flies Recaptured
Daily Mean' Trap Mean
Total (flies/day) (flies/trap/day)
44 6.3 0.3
58 8.3 0.4
3 0.5 0.0
1 0.1 0.0
0 0.0 0.0
106 3.0 0.2
38 5.4 0.3
7 0.5 0.0
3 0.4 0.0
81 11.6 0.6
9 1.3 0.1
6 0.9 0.0
144 3.3 0.2
250


All weeks except weeks 3, 4 (6 d) and wk 5 (21 d) consisted of 7 d collection periods. Number of traps placed weekly: 2008, 20 traps;
2009, 18 traps.
a Total house flies captured includes first-time capture of unmarked house flies and recapture of marked and released house flies.
b Flies collected in wk 4 and 7 were recaptured from previous releases in wk 3 and 6 respectively, indicating more than 7 d longevity
in the field. These data were not used in any analysis.
' Daily means (no. flies/d) for each test period were calculated by dividing total capture numbers by the duration (d) of that period.
d Trap means (no. flies/trap/d) were calculated by dividing daily mean house fly capture numbers by the number of traps in use, i.e.,
wk 1-5, 20 traps; for wk 6-11, 18 traps.











Table 2-5. House fly release week and recapture rate and associated weekly weather data obtained from Weather Underground station
approximately 30 km from the release site in north central Florida.
Temperature (C) Precipitation Wind Speed Wind Dir Barometric Pressure (hPA)
(km/h)

Wk Recapture Max Min Mean Total (cm) Mean Max Prevailing Min Max Mean
Rate (%)

1 0.08 30.1 19.4 24.8 0.38 9.7 24 ENE 1006 1033 1019
2 1.93 26.3 12.6 19.5 0.04 9.7 24 ENE 1006 1035 1020
3 0.60 22.1 7.2 14.8 0.24 9.7 24 WSW 1007 1033 1020
4 0.20 22.1 7.2 14.8 0.46 9.7 24 WSW 1013 1018 1020
5 0.00 25.0 9.5 17.5 0.76 8.1 24 SSE 1001 1031 1016
6 0.63 20.6 5.7 12.9 0.30 8.1 27 WSW 1013 1023 1018
7 0.12 25.2 17.0 21.3 0.46 12.9 35 ENE 1009 1023 1016
8 0.15 29.5 19.4 24.7 0.46 8.1 27 SSE 1009 1023 1016
9 0.81 31.3 19.8 25.7 0.33 8.1 27 SSW 1011 1015 1016
10 0.15 30.6 20.5 25.6 0.57 6.4 29 WSW 1011 1016 1013
11 0.15 33.4 21.7 27.7 0.63 8.1 30 SSE 1005 1016 1013


Wk 1-5 (23 October 2008 to 4 December 2008) and wk 6-11 (21 May 2009 to 25 June 2009).









CHAPTER 3
ESCHERICHIA COLI0157:H7 PREVALENCE

Introduction

Escherichia coli 0157:H7 is a bacterial pathogen that causes dysentery and

diarrheagenic diseases in humans, but is commensally present in its primary reservoir,

cattle. Escherichia coli 0157:H7 has been isolated from house flies, Musca domestic L.,

on dairies (Alam and Zurek 2004). Florida has a large number of dairies that are located

in close proximity to human population centers, and the expansion of many urban areas

has decreased the distance between dairies and towns. House flies typically disperse

within 3.3 km (Parker 1916) to 5 km (Peppler 1944), although they move freely within

urban areas for up to 6.7 km (Quarterman et al 1954). Additionally, house flies are

capable of flying as far as 8 km from their breeding sites (Bishopp and Laake 1921). The

ability of house flies to disperse within and between rural and urban areas indicates that

house flies have tremendous potential to serve as vectors for transmitting E. coli

0157:H7 from the dairies to the town centers, particularly to restaurants. The close

proximity of dairies to towns might permit the pathogen to cause human disease by the

fecal-oral route through involvement of the house fly. Although E. coli 0157:H7 has

been isolated from dumpsters outside restaurants in Gainesville, FL (Butler et al. 2010),

no study has been done in Florida to determine the potential of pathogen transmission

from dairies into towns by house flies.

Diarrheal diseases impart an enormous toll on human and agricultural animal

(e.g., cattle) populations, with severe health and economic impacts. Hospital expenses for

human patients with infectious diarrhea can be four times greater than those of other

patients. Similarly, medication expenses can be four times higher and the length of









hospitalization can be three times longer (Suda et al. 2003). Economic impacts include

lost income for family members who must miss work, as well as lost profits for

employers (Buzby et al. 1996, Abe et al. 2002, Sapers and Doyle 2009).

For cattle, increased operating expenses are incurred by dairy farms, feedlots and

cattle rendering plants, sanitation efforts, including fly surveillance and management

measures to comply with food-safety federal regulatory mandates (CDC 2009). Lawsuits

can result in imposition of stiff fines upon many different food-related industries

including producers, distributors, and restaurants. A recent example: in June, 2008, a

lawsuit led to a $13.5 million settlement after a child in Milwaukee, Wisconsin died due

to consumption ofE. coli 0157:H7-contaminated food (Powell 2008, Rohde, 2008).

Enteric bacteria such as E. coli 0157:H7 can be transmitted by the house fly, M.

domestic, and other flies (Greenberg 1971, Greenberg 1973). Cattle are a primary

reservoir for E. coli 0157:H7 (Heuvelink 2003), and dairy farms located near towns or

residential areas may present a potential public health threat if pathogen persistence and

high populations of filth flies co-occur and if flies were to disperse from the dairy. House

flies typically disperse no further than 0.3-1.2 km (West 1951, Quarterman et al. 1954,

1964, Stein 1986, Milio et al. 1988, Alam and Zurek 2004), although they can travel as

far as 8 km (13 mi) from their breeding sites (Bishopp and Laake 1921). In a recent study

at a dairy in north-central Florida, I determined that house flies dispersed up to 3 km into

a nearby town (Chapter 3). These characteristics of house fly behavior increase the

potential for the introduction of pathogens from cattle to humans, particularly from sites

conducive to fly-breeding (e.g., dairies) (Kaufman et al. 2005b) to nearby human

population centers.









Isolation and identification ofE. coli 0157:H7 by standard microbiological

methods is time-consuming and labor-intensive. Because infection with E. coli 0157:H7

is a nationally notifiable event in the United States (Mead and Griffin 1998), detection

methods that are rapid, selective and sensitive are important (Ogden et al. 2001).

Szalanski et al. (2004) developed a 6-h protocol for detection of E. coli 0157:H7 from

house flies by polymerase chain reaction (PCR) instead of standard microbiological

techniques. Many researchers use PCR to definitively confirm presence of virulence

factor genes after presumptively confirming species identification by means of multiple

microbiological and biochemical tests (Buma et al. 1999, Cagney et al. 2004, Alam and

Zurek 2004). Usage of PCR techniques following microbiological methods improves

identification and characterization of E. coli 0157:H7 because more than 100 E. coli

serotypes produce Shiga-like toxins (Cebula et al. 1995). Presumptive identification of E.

coli 0157:H7 by microbiological methods can eliminate many serotypes prior to PCR

analysis, and PCR can subsequently be used to confirm identification and to serotype

additional characteristics of E. coli 0157:H7. Use of PCR in place of further biochemical

tests can save both time and money in the laboratory, while increasing isolation

sensitivity.

The pathogenicity of enterohemorrhagic E. coli 0157:H7 is dependent upon

possession of several virulence factors that are encoded in the genome or in a large

plasmid. PCR provides a more selective and sensitive method to identify E. coli 0157:H7

in samples (Visetsripong et al. 2007), and permits serotype differentiation (Beutin et al.

2007), and enables a faster identification of E. coli 0157:H7 than direct culture methods,

which is especially important during outbreaks. Primer pairs can be used in uniplex or









combined in multiplex PCR to detect and amplify target gene fragments of specific

virulence factor genes, such as the rfbEH7 andfliCois7 genes.

In this study the prevalence of E. coli 0157:H7 in house fly, grain, and manure

samples from two dairies and in house fly samples from two restaurant garbage

dumpsters in a nearby town was determined by direct culture and confirmed by PCR

detection of the rfbEH7 andfliCois7 genes.

Materials and Methods

The overall study area was the same as that described previously (Chapter 2).

House fly populations were monitored at the feed barns of two dairies, A and B, with

Dairy B located 1.5 km east of Dairy A. Samples were collected for microbiological

analysis from four sites: both dairies and two restaurants, C and D, located 3 km and 3.5

km southeast of Dairy A, respectively. Collection sites were chosen following selection

of a town in north-central Florida that met all of the following criteria: 1) one or more

dairies located within 3 km of the town; 2) one or more restaurants in town with a

dumpster located adjacent to the restaurant; and, 3) a nearby residential human

population. This research model represents a reasonable set of naturally-existing

conditions under which a potential for pathogen transmission from dairies to restaurants

exists. These conditions allowed E. coli strains collected at dairies and restaurants to be

compared.

Three laboratories in Gainesville, Florida, were used during this study. The Food

and Environmental Toxicology (FET) Laboratory at the University of Florida was used to

isolate and identify E. coli 0157:H7 from grain, manure, and pooled house fly samples

using direct culture methods. The Veterinary Entomology (VE) Laboratory at the

University of Florida was used to store untested samples for future analysis by









polymerase chain reaction. The United States Department of Agriculture, Agricultural

Research Service, Center for Medical, Agricultural and Veterinary Entomology (USDA-

ARS-CMAVE) laboratory was used to measure house fly population counts as described

below.

House fly population trends were monitored weekly during selected intervals at

both dairies from 7 June 2008 to 23 September 2008 using sweep nets (i.e., small

"butterfly" nets), Scudder grids (Scudder 1947), a portable slatted wood frame placed on

top of fly breeding areas, and fly spot cards (Lysyk and Axtell 1985), 7.5 x 12.5 cm index

cards placed on walls on which flies excrete and regurgitate while resting (see Chapter 2

for a full description). Use of multiple monitoring methods was performed with the

intention of examining these data for possible trends between these three monitoring

methods (Table 3-1) as well as for correlation between house fly populations and E. coli

0157:H7 prevalence. Nets, grids and cards were all used as close to fly aggregation and

breeding areas as possible, and collection locations were the same every week unless

noted differently.

"Snapshot" active monitoring of house fly populations by sweep nets and Scudder

grids was discontinued after 23 September 2008 with the cessation of microbiological

sampling, but weekly spot card sampling was continued until 4 December 2008. Because

spot card data may represent activity of multiple species of flies, sticky cards, index cards

(7.5 cm x 12.5 cm) covered with adhesive on one side so that flies become captured when

they land (see Chapter 2 for a full description), were placed along barn walls at Dairy A

from 30 October 2008 to 4 December 2008. Sticky cards permitted differentiation

between fly species. Thus the sticky card and spot card data were examined for potential









trends or correlations of house fly populations on Dairy A, the release site in my dispersal

study (Chapter 2). Alsynite traps were also placed at each dairy 1-2 m outside of the feed

barns at each dairy from 16 September 2008 to 4 December 2008 as part of a house fly

dispersal study (Chapter 2). Because some of the alsynite traps were in use at the feed

barns when spot and sticky cards were in use, the alsynite traps provided an additional

method of house fly population monitoring and allowed for examination of correlations

between monitoring methods. House fly monitoring occurred again from 14 May 2009 to

25 September 2009 using spot cards, sticky cards and alsynite traps at Dairy A, and spot

cards and alsynite traps at Dairy B.

Sweep net monitoring of adult house flies was performed at both dairies by

sweeping an insect net (45-cm diameter, Mod. No. 7112NA, Bioquip, Rancho

Dominguez, CA) in one figure-eight pattern. Sweeps were performed so that the net was

at a height of 0.5 m above adult aggregation areas in feed trough areas. Sweep nets were

also performed above spilled grains that were below feed augers. Sweep nets were

performed while walking towards the sun, to prevent disturbance of house flies due to

casting a shadow over resting flies and at a fast walking pace. One to four net sweeps

were performed at each dairy from 7 June 2008 to 23 September 2008. Mean sweep net

counts for each dairy were calculated for each collection day by dividing the total number

of flies captured by the number of sweeps performed.

Small Scudder grids (45 x 45 cm) (Scudder 1947, Murvosh and Thaggard 1966)

(Fig. 3-1) consisted of 12 slats of rough unfinished wood (8 mm x 45 cm) laid parallel to

each other and spaced evenly along a 45 x 45 cm wood frame. This provided a series of

alternating edges for flies to rest upon. Scudder grids were placed on the ground on top of









spilled grains under augers or on top of feed troughs 1-10 times at each dairy from 7 June

2008 to 23 September 2008. Each Scudder grid was photographed after it was in place for

a period of 5 sec, and the number of house flies on each grid was counted by examining

digital photographs on a computer monitor at the USDA-ARS-CMAVE laboratory.

Where photographs were not available, estimates were noted on the data sheet.

Ten spot cards were placed at each dairy inside a barn adjacent to the milking

parlor (Dairy A, milk barn; Dairy B, south barn) on 5 August 2008. Because use of the

milk barn at Dairy A discontinued in September 2008, spot cards were relocated to Dairy

A's feed barn located approximately 0.25 km north of the milking parlor on 23 September

2008. Use of spot cards was continued in Dairy A's milk barn until 30 October 2008 to

track fly population changes after departure of the cattle. Spot card monitoring at each

dairy using Dairy A's feed barn and Dairy B's south barn continued until 4 December

2008, and was resumed from 14 May 2009 to 25 June 2009. Spot cards were placed

horizontally at a height of 2-3 m and were flush against either support beams or rafters. In

all barns, spot cards were placed at an approximate height of 2-3 m and in locations

where they were not within reach of the cattle. Spot cards at Dairy B and at Dairy A's

milk barn were inserted into metal frames nailed to the wooden support beams of these

barns and held in place with one small binder clip placed at the top edge of the metal

frames (Fig. 3-2), or by clipping them to large metal support beams with small and

medium binder clips (Office Depot, Delray Beach, FL) (Fig. 3-3).

Sticky cards (7.5 cm x 12.5 cm) (Hogsette et al. 1993) were used from 30 October

2008 to 4 December 2008, and again from 14 May 2009 to 25 June 2009. Fourteen sticky

cards were placed at Dairy A's feed barn by clipping them with large paper clips to the









steel support beams immediately below the spot cards so a gap of approximately 1 cm

existed between the spot and sticky cards (Fig. 3-3).

Three alsynite sticky sleeve traps (Broce 1988, Hogsette and Ruff 1990) were

used for 5 wk from 30 October 2008 to 4 December 2008 in conjunction with my 2008

dispersal study (Chapter 2). Two traps were located at Dairy A and one was located at

Dairy B (Chapter 2, Fig. 2-1). All alsynite traps were placed within 1-2 m of a specific

barn or feed trough as described below. The traps at Dairy A were placed 100 m apart,

with one trap (trap 1) at the SE corer and the other at the SW corner of Dairy A's feed

barn (trap 2). The trap at Dairy B was placed at the SE corner of the south barn (trap 8).

Five traps were used at the dairies from 14 May 2009 to 25 June 2009 in conjunction

with my 2009 dispersal study (Chapter 2), with two at Dairy A (traps 1 and 2) and three

at Dairy B (traps 8, 24 and 25). Trap locations used in 2008 were used again in 2009.

Two additional alsynite traps were placed on Dairy B during 2009: one (trap 24) was

located W of the calf feed trough, approximately 0.3 km S of the adult barns, and the

second (trap 25) was placed at the NE corner of the adult barns approximately 100 m

from the original alsynite trap location used in 2008. These alsynite traps were relevant to

fly monitoring efforts because they were placed immediately outside of barns on both

dairies where cattle are fed and where adult flies aggregate.

In addition to house fly population monitoring, I collected samples to test for the

presence of E. coli 0157:H7 by direct culture microbiological analysis. Samples

collected from dairies included adult house flies, spilled grains, and fresh manure. At

restaurants, only adult house flies from the garbage dumpsters were sampled.









Flies were collected by sweep-netting, with the additional use of sterilized

materials. All materials that came into direct contact with flies were either purchased

sterile, autoclaved for 60 min at 121 OC, 15 psi, or placed in 6% hypochlorite (bleach)

(Clorox Co., Oakland, CA) solution for 15 min prior to use. Sweep nets were autoclaved

and remained sealed in autoclave bags until used to collect a single sample in the field.

Each sample of flies collected by sweep netting was transferred to a sterile 120-ml clear

polypropylene specimen cup (Model 70756, Samco Scientific Corp., San Fernando, CA).

Fresh nitrile gloves (Best Glove, Inc., Menlo, GA), sterilized with 6% bleach (Clorox

Co., Oakland, CA), were worn when using each sweep net.

Specimen cups containing fly samples were placed on ice, in a large cooler (50 L,

Igloo Corp., Houston, Texas) for approximately 1 min (10-20 C), to chill flies without

mortality. Up to 10 flies were gently shaken onto a chilled metal pan (lined with

aluminum foil which was changed after every sample) and identified to species. Flies not

removed from specimen cups for sorting were later identified in the laboratory. Up to 10

M. domestic were placed individually into sterilized snap-cap microcentrifuge tubes

(P/N 02-681-240, Fisher Scientific Co., Waltham, MA). Flies were manipulated using

sterilized feather-weight forceps (P/N 4750, Bioquip, Rancho Dominguez, CA). The 10

individually contained flies were placed inside a pre-labeled sealable, clear, plastic bag,

and placed in the cooler. All fly samples remained in the cooler at 10-20 C for 2-6 h

until removed for laboratory processing later that same day.

During the cooler months of November-April when fly populations were low,

isolation of 10 individual flies was given priority over pooled samples; i.e., if only two

house flies were collected, then each of the two was placed individually in sterile 1.5-ml









cap microcentrifuge tubes. If 12 flies were collected, then ten were individually placed in

microcentrifuge tubes and the remaining two flies were retained in the specimen cup.

During the warmer months of May-October, flies collected at each site were transferred

to sterile specimen cups, placed on ice in a cooler, transported to the laboratory, and

knocked down by placement in -20 C for approximately 1 min. Afterwards, specimen

cups were removed from the freezer, and 10 individual flies were transferred into

individual microcentrifuge tubes as described previously.

Cattle manure was obtained using sterile materials. Manure samples consisted of

fresh droppings, and were obtained within 30 sec after animal defecation. Slurry samples

were collected from barn floors or lagoons (wastewater retention ponds). Grain samples

were obtained from feed troughs or from spilled grains below feed augers.

Manure, and grain (substrate) samples were obtained manually by two methods

(Fig. 3-4). In the first sampling method, approximately 100 g of each substrate was

scooped out with a sterilized metal spoon. Manure samples were obtained from the center

surface of the fresh dropping. Manure samples were placed in pre-labeled 120-ml sterile

specimen cups and placed inside the cooler (10-20 C) for transport to the laboratory

where they were processed within 24 h of collection. In the second sampling method, two

sterile swabs, pre-moistened with buffered peptone water (BPW) (Oxoid Ltd.,

Basingstoke, Hampshire, England) or trypticase soy broth (TSB)(Difco, Becton,

Dickinson and Co., Sparks, MD), were inserted 2.5-5.0 cm deep into manure or grain,

and rolled around briefly until completely coated with substrate particulates. Both swabs

from each substrate were placed together as one sample in sterile 15-ml polypropylene

centrifuge tubes (Model No. 35-2096, Becton Dickinson Labware, Franklin Lakes, NJ)









containing 9 ml of sterile BPW or TSB. Swab samples were placed in a shaded location

in the vehicle, and transported to the laboratory at ambient temperature (25-30 C).

Air temperature, relative humidity (RH), and barometric pressure (BP) data were

recorded on-site at Dairy A using a portable weather station (Model No. 00589W, Acu-

Rite, Jamestown, NY) and examined to determine if any of these weather conditions

influenced house fly population activity determined by house fly monitoring methods.

At the time of sample collection, both the substrate and surface temperatures of

manure and grain samples were measured. Substrate temperatures were measured by

inserting a digital soil probe thermometer (Model No. 6310, Spectrum Technologies, Inc.,

Plainfield, IL) 2.5-5.0 cm into the grain or manure. The digital thermometer was allowed

to calibrate while samples were obtained. Surface temperatures were measured by

pointing an infrared thermometer (Raynger ST2, Raytek Corp., Santa Cruz, CA) at the

center of the sample from a distance of 1 m.

All samples that were scheduled to be processed while fresh using standard

microbiological methods were delivered to a Biosafety Level 2 (BSL2) laboratory in the

FET Laboratory within 6 h of collection, and processed within 24 h of collection. Fly

samples were placed in a -20 C freezer for up to 30 min to reduce the potential for flies

escaping during processing; substrate samples remained in the cooler at 10-20 C, and

swab samples were maintained at room temperature (25 C) until processed. Some

samples were reserved by storage in a -20 C freezer at the VE Laboratory for future

identification and serotyping by polymerase chain reaction (PCR).

All samples were processed using standard BSL2 laboratory aseptic techniques,

and in compliance with the University of Florida's (UF) Environmental Health and Safety









(EHS) Biosafety Protocols (UF-EHS 2008). Pipette tips, tubes, broths, and other reagents

were either purchased sterile or were autoclaved for 60 min at 121 OC, 15 psi, except

where noted differently, using a table-top autoclave (Sterilmatic STME, Market Forge,

Ramsey, MN). Agar plates were poured and allowed to solidify inside a BSL2 cabinet.

Samples were processed at an open BSL2 bench as described below.

Aerobic plate counts (APCs) of unenriched background microbial (BM)

organisms were performed. Each sample was vortexed for 30-60 sec to resuspend

bacteria and held at room temperature for 1 min to permit debris to settle. A 1-ml aliquot

of each unenriched sample was pipetted into a 15-ml (O.D. x Length 16x125 mm)

borosilicate glass screw-capped culture tube (No. 9825-16X, Corning Pyrex Inc., Lowell,

MA) containing 9 ml of BPW.

Serial dilutions were prepared for each sample by sequential pipetting 1 ml of the

bacteria:broth mixture to tubes containing 9 ml BPW, for a total of six serial dilutions,

10- 10-6. Two aerobic plate counts using Petrifilm APC plates (3M, St. Paul, MN) were

performed for each dilution tube. First, tubes were vortexed for 30-60 sec and allowed to

settle for 5-10 sec. Then 1-ml aliquots were pipetted onto APC plates. In this way, the

resultant culture plate dilutions ranged from 10-1 10-6 colony forming units per gram

(CFU/g).

A new pipette tip was used for each tube, and dilutions were transferred

sequentially from the most to the least dilute, to decrease the possibility of cross-

contamination. After the 1-ml aliquot of each dilution was placed on an APC plate, a

proprietary plastic tool (spreader) that was included with the PetrifilmTM APC plates was

gently placed upon the APC plate to spread the diluted sample out until it covered a 20-









cm2 surface. Plates were incubated at 37 C for 6-18 h and plate counts were performed

the following day by counting CFUs on each plate. Plates with 14-300 colonies were

used to calculate average values for total CFU/g of aerobic bacteria for each sample.

To improve selectivity for E. coli 0157:H7 from complex fecal and decomposing

organic matter containing high concentrations of competing background microorganisms,

TSB was modified by the addition of novobiocin (20 mg/1) (mTSB+N) (FDA-CFSAN

2007a) for the enrichment broth (Desmarchelier et al. 1998).

Bacteria species used as positive and negative controls in this study were obtained

from Dr. Huang (Auburn University, AL) and are maintained by Dr. Simonne in the FET

Laboratory. Nalidixic acid-resistant E. coli 0157:H7 strain 204P was used as a positive

control. Negative controls used in this study were .\iigell,/ dysenteriae (ATCC 49550)

and Salmonella ilh,,nl,,ui (ATCC 8391).

Frozen fly samples containing multiple fly species were removed from the

freezer, and up to 25 M. domestic were randomly selected and placed as a pooled

sample in mTSB+N. Species other than M domestic were discarded. Adult house flies

were incubated individually or in pools of up to 25 flies. Some thawing occurred during

sorting. For pooled samples with up to nine flies, M. domestic adults were added to 9 ml

mTSB+N in a sterile 15-ml polypropylene centrifuge tube (Model No. 35-2096, Becton

Dickinson Labware, Franklin Lakes, NJ). Pools containing 10 or more adult flies were

placed in a 50-ml polypropylene centrifuge tube (Model No. 43089, Corning

Incorporated, Coming, NY) containing 25 ml mTSB+N. Individual samples consisted of

one M. domestic adult that was placed into a sterile 15-ml polypropylene centrifuge tube









(Model No. 35-2096, Becton Dickinson Labware, Franklin Lakes, NJ) containing 9 ml

mTSB+N. Samples were incubated at 37 C for 24+2 h.

Substrate samples were initially enriched using 25 g of each substrate. However,

following challenges with odor in the FET Laboratory, a swab sample technique was

adopted (Rice et al. 2003, Greenquist et al. 2005, Davis 2006). In the initial substrate

enrichment process, 25 g of each substrate sample were added to individual double-

bagged 400-ml stomacher bags (177 mm x 305 mm) (Stomacher 400 Classic, P/N

BA6041/CLR, Seward Co., Seward, UK) containing 225 ml of mTSB+N. The substrate-

mTSB+N mixture was homogenized in a stomacher for 60 sec. Unused portions of

substrate samples from each bag were stored in a freezer at -20 oC for future analysis.

Some of the unused substrate samples were disposed of following mechanical failure of

the freezer that resulted in thawing of samples. In the second substrate enrichment

process, swab sample tubes (Model No. 35-2096, Becton Dickinson Labware, Franklin

Lakes, NJ) containing 2 substrate-inoculated swabs in 9 ml BPW were processed by

vortexing each tube for 30-60 sec. One swab and 1 ml supernatant was transferred to a

new 15-ml centrifuge tube (Model No. 35-2096, Becton Dickinson Labware, Franklin

Lakes, NJ) containing 9 ml of mTSB+N. Transferred samples were incubated at 37 C for

24+2 h. The second swab and remaining BPW supernatant (8 ml) were stored unaltered

in a freezer at -20 C for future analysis. Following selective enrichment, enriched

samples were removed from the incubator, vortexed for 30-60 sec and allowed to settle

for 1 min in preparation for enumeration and isolation procedures.

Enumeration ofE. coli 0157:H7 was performed using SMAC plates

supplemented with cefixime (15 [tg/l) and potassium tellurite (1.25 [tg/l) (CT-SMAC)









(Alam and Zurek 2004, FDA-CFSAN 2007b). One-ml aliquots of each unenriched

vortexed sample were transferred to screw-capped culture tubes containing 9 ml of sterile

BPW and vortexed for 30-60 sec. Six serial dilutions were prepared for each sample, as

previously described, and 100 pl aliquots were pipetted onto each of two CT-SMAC

plates, i.e., double-plated, so that two plates were prepared for plate dilutions that ranged

from 10-2 to 10-7. The unenriched bacteria-broth mixture was spread evenly over the plate

using a glass rod. Spread-plate counts ofE. coli 0157:H7 were obtained by averaging the

number of CFUs for plates that contained 14-300 colonies. CT-SMAC spread-plates were

incubated at 37 C for 24+2 h.

Isolation ofE. coli 0157:H7 was performed by immunomagnetic separation

(IMS) and direct culture, as described below. Separation, following Invitrogen's

proprietary protocol (Dynal 2007), was performed using magnetic E. coli 0157-specific

antibody-coated beads (Dynabeads anti-E. coli 0157, Invitrogen, Carlsbad, CA) that

increased isolation sensitivity. Resuspended beads (20 al, [108 beads/ml]) were aliquotted

into a 1.5-ml microcentrifuge tubes and 1 ml of enriched house fly sample supernatant

was added to each tube. Tubes were placed in a manually-operated magnetic stand,

(Ambion 6 Tube Magnetic Stand, Invitrogen, Carlsbad, CA), closed and inverted gently

for 10 min at room temperature (RT) to permit the E. coli 0157:H7 to bind antigenically

to the beads while keeping the beads suspended in the supernatant.

A magnetic plate was inserted into the magnetic stand, and tubes were inverted

multiple times to form a pellet of concentrated beads at the bottom of the tube, along the

magnetized side of the tubes. Tubes were allowed to stand for an additional 3 min at RT

to maximize recovery of beads coated with E. coli 0157. Supernatant was pipetted out









and discarded, taking care not to dislodge the bacteria-bead pellet complex. The magnetic

plate was removed and three sequential 1-ml 20X PBS Tween-20 (Thermo Scientific,

Rockford, IL) wash buffer rinses were performed to resuspend the beads and enhance

removal of non-specific binding microorganisms.

After the third rinse, the bacteria-bead complex was resuspended in 100 [l of PBS

Tween buffer and vortexed. The entire 100 [l bacteria-bead complex was plated onto a

CHROMAgar (BBLTM CHROMAgarTM 0157, Beckton Dickinson Co., Franklin Lakes,

NJ) plate by moistening a sterile swab with the bacteria-bead complex and streak-plating

half the agar plate with the moistened swab. The second half of the plate was loop-

streaked to enhance isolation of colonies. Plates were incubated at 37 C for 24+2 h.

After incubation, both CT-SMAC spread-plates and CHROMAgar isolation plates

were examined for presumptive E. coli 0157:H7 colony growth. Presumptive E. coli

0157:H7 appeared colorless with or without a light smoky center (sorbitol-negative) on

CT-SMAC plates, and light-violet to violet on CHROMAgar plates. Up to five

presumptive-positive E. coli 0157:H7 colonies from CHROMAgar plates were selected

and streaked onto CT-SMAC plates which were incubated at 37C for 242 h. Presumed

E. coli 0157:H7 colonies were subsequently loop-streaked onto non-selective, general

growth Trypticase soy agar with yeast extract (TSAYE) plates and incubated at 370C for

24+2 h.

Sub-cultures were prepared for long-term storage by transferring a loopful of one

TSAYE colony into 850 [l of a sterile Luria-Bertani (LB) broth. The bacteria:broth

mixture was pipetted into 150 sterile [il glycerol in 2-ml cryogenic vials (P/N 10-500-26,

Fisher Scientific, Pittsburgh, PA). The LB broth, bacteria, and glycerol were gently









pipetted in and out until thoroughly mixed (5-10 times) without introduction of air

bubbles. Finally, the bacterium and its 15:85 glycerol:bacteria-broth mixture were

vortexed for 30 s and stored at -20 oC, to be available for further DNA analysis by

polymerase chain reaction (Chapter 5).

Where sufficient samples were available, additional biochemical tests were

conducted. These tests included phenotypic expression of characters typical for E. coli

0157:H7: 1) hydrolization of tryptophan; 2) the presence of a brilliant green sheen on L-

EMB agar; and, 3) a lack of fluorescing 4-methylumbelliferyl-P-D-glucuronide (MUG).

Hydrolization of tryptophan was assessed by placing a filter paper wetted with

Kovac's Reagent (Ricca Chemical Co., Arlington, TX) on each positive-growth plate.

Colonies that are presumptive positive for E. coli 0157:H7 typically turn pink.

Presumptive E. coli 0157:H7 colonies were plated concurrently onto two different

media; Levine's eosin methylene blue (L-EMB) (Oxoid) agar, and fresh TSAYE agar

plates. Incubation of L-EMB plates, which inhibit gram-positive growth, occurred at 37

C for 242 h. Following incubation, plates were examined for a distinctive bright

metallic green sheen on dark-blue to black nucleated colonies. This brilliant green sheen

is characteristic of E. coli 0157:H7, and differentiates E. coli 0157:H7 from non-

pathogenic E. coli that turn dark green without the brilliant green sheen on L-EMB agar.

Isolates that were transferred to TSAYE plates were tested for the presence of

glucoronidase, an enzyme that hydrolyzes 4-methylumbelliferyl-P-D-glucuronide (MUG)

to yield fluorescing 4-methylumbelliferone, by addition of one ColiComplete (Bothell,

WA) disk to the plate's heaviest bacterial streak. After incubation at 37 C for 242 h,









colonies presumptive for E. coli 0157:H7 were considered negative if they lacked blue

fluorescence around the disks when examined under 365 nm ultraviolet (UV) light.

Enumeration of aerobic bacteria was performed using Petrifilm APC plates

containing 14-300 CFU/g. Aerobic bacteria counts were obtained for 19 selected samples

that consisted of nine house fly, seven grain and three manure samples that were

collected from 31 May 2008 to 26 August 2008. Aerobic plate counts of the two grain

and two house fly samples obtained from Dairy A on 26 August 2008 were averaged for

each type. Enumeration data for the remaining 17 samples were based on single samples.

When multiple dilution plates containing 14-300 CFU/g were obtained from individual

samples, the range of those plates was recorded.

Presumptive E. coli 0157:H7 colonies were counted using CT-SMAC spread plates

containing 14-300 CFUs on two dates: 31 May 2010 and 14 June 2010. Samples of 31

May 2010 were enumerated, but not processed further. Samples of 14 June 2010 were

submitted for microbiological analysis after enumeration ofE. coli 0157:H7 was

completed.

Colonies that tested positive on CHROMAgar plates were classified as presumptive

positive. However, due to failure of the CHROMAgar on 26 August 2008 and 16

September 2008, isolates that were presumptive positive on CT-SMAC agar plates on 26

August 2008 and 16 September 2008 were also submitted to PCR. Thus, isolates of 24

samples of the 57 dairy-collected samples were submitted to PCR, with isolates from 11

samples originating from CHROMAgar plates and isolates from 13 samples originating

from CT-SMAC plates.









Multiple isolates were obtained from several samples due to sequential transfer of

individual colonies during the microbiological testing. Therefore, prevalence rates were

determined for both the number of samples tested and for the number of isolates that

were sub-cultured from samples.

Sub-cultures of each isolate were prepared for long-term storage by aseptic loop-

transfer of one loop of logarithmic-growth phase E. coli 0157:H7 from TSAYE culture

plates to 850 ptl sterile LB (Oxoid) broth (lysogeny broth; Bertani (1951, 2004)). The LB

broth and bacteria were mixed by gentle pipetting and then added to 150 ptl sterile

glycerol in a sterile 2-ml cryogenic vial (P/N 10-500-26, Fisher Scientific, Pittsburgh,

PA). The LB broth, bacteria, and glycerol mixtures were stored as a 15% glycerol stock

in -20 C non-thawing freezer. Presumptive positive isolates were prepared for DNA

extraction and PCR analysis from 1 December 2009 to 16 April 2010, approximately 1.5

yr after placement in the freezer.

To prepare frozen isolates for DNA extraction and PCR, fresh cultures of each

isolate were prepared from glycerol stock by transferring 10 tl of bacteria into a 15-ml

culture tube containing 2 ml sterile LB broth. Transfer of bacterial stock was performed

in a BSL2 cabinet. The stock was returned to the freezer to minimize thawing. Culture

tubes containing fresh bacteria-broth mixtures were incubated at 37 C for 24+2 h with

shaking (250 RPM) (Lab-Line Orbit Environ-Shaker, Lab-Line Instruments Inc.,

Melrose Park, IL). Eight hundred fifty microliters of this fresh culture were used to make

another glycerol stock to be placed in a -70 C freezer for long-term storage, and the

remaining culture was used for DNA extraction and PCR amplification.









Total genomic DNA was extracted from stored bacterial isolates originally

obtained from house fly, grain and manure samples using the QIAquick DNeasy Blood

and Tissue Kit (QIAgen, Valencia, CA). Extracted DNA was resuspended in 100 [l

Buffer AE and stored at 4 OC.

Polymerase chain reaction was performed using one multiplex and two uniplex

PCR assays (Table 3-2) to amplify gene fragments using two primer pairs (20 pmol/ml

each) designed to target the rfbEH7 andfliCols5 genes (Hu et al. 1999, Cagney et al. 2004,

Szalanski et al. 2004). ThefliCo157 primer pair amplifies a 625-bp E. coli 0157 serotype

gene fragment (Gannon et al. 1997) and the rfbEH7 primer pair amplifies a 259-bp E. coli

H7 gene fragment (Paton and Paton 1998).

Following low amplification of both gene fragments simultaneously in the

multiplex PCR, samples were submitted to uniplex PCR for each gene fragment

separately. The following three assays were conducted to test for the genes of interest:

Assay 1, Multiplex PCR for rfbEoi57 andfliCH7; Assay 2, Uniplex PCR for rJbEoi57; and

Assay 3, Uniplex PCR forfliCH7.

Master mix reagents for each assay consisted of deionized sterile water, 10X

DNA Polymerase PCR Buffer (-MgC12), dNTP Mix (10 mM each), 50 mM MgCl2,

oligonucleotide primers (20 pmol/pl each) (Eurofins MWG Operon, Huntsville, AL)

(Table 3-2), and recombinant DNA polymerase Taq enzyme (Invitrogen, Carlsbad, CA).

For the multiplex assay and for the amplification of the rfbEH7, the PCR program

consisted of an initial denaturing at 94 OC for 2 min, followed by 35 cycles of denaturing

at 94 C for 45 sec, annealing at 56 C for 45 sec, and extending at 72 C for 1 min. A

final extension was performed at 72 C for 5 min. The PCR program for amplifying









flicCoi57 gene region was similar to those just described but the annealing temperature

was increased to 65 C.

Sterile distilled water was used as a negative control in each assay. Bacterial

positive and negative controls used in this study were obtained from Dr. Amarat Simonne

at the University of Florida, in the Food and Environmental Toxicology Laboratory.

Nalidixic acid-resistant E. coli 0157:H7 strain 204P was used as a positive control.

Negative controls were .\iigel.// dysenteriae (American Type Culture Collection (ATCC)

49550) and Salmonella iih,,q,,nui (ATCC 8391). Cultures used for this study were

maintained in the laboratory of Dr. James E. Maruniak at the University of Florida,

Entomology and Nematology Department, with subcultures stored at both -20 C and at -

70 oC.

Amplified PCR gene fragments were visualized by ethidium bromide staining of

the PCR product that was separated in 1% agarose gel electrophoresis. Visualized PCR

products were photographed using ultraviolet (UV) light. Isolates were considered

positive if both thefliC and rfbE gene fragments were amplified. Because multiple

isolates corresponded to each dairy-collected sample, samples were considered positive if

at least one isolate was positive. The percentage of positive samples was calculated by

dividing the number of positive samples by the number of samples that were submitted to

PCR and multiplying by 100.

DNA Quantification

DNA concentration of freshly cultured isolates was obtained by measuring the

absorbance of light at wavelengths of 260-280 nm, using a spectrophotometer (Nanodrop

1000 mini-spectrophotometer, Thermo Scientific, Waltham, MA) or was estimated by

visual comparison of the band intensity with the standard 100 bp ladder purchased from









Invitrogen (Genvault 2010). Spectrophotometry nanodrop analysis was performed in

accordance with the manufacturer's instructions in Dr. Michael Scharfs laboratory,

Entomology and Nematology Department, University of Florida.

16S rDNA PCR Analysis

To ensure that PCR-product gene fragments were from bacterial DNA, a broad-

range 16S rDNA PCR assay was performed on 46 selected isolates. Broad-range 16S

rDNA PCR analysis was performed in Dr. Volker Mai's laboratory, Emerging Pathogens

Institute, University of Florida.

Statistical Analysis

Seasonal trends of house fly populations were estimated for the sampling period

using weekly mean counts for both passive and active fly monitoring methods at both

dairies during the respective weeks of placement of each passive monitoring method.

Passive fly monitoring estimates were obtained with spot cards, sticky cards and sticky

sleeves on alsynite traps. Active monitoring estimates were obtained with weekly active

"snapshot" counts of Scudder grids and sweep nets at four sampling sites.

House fly population data were subjected to PROC UNIVARIATE to examine

normality and PROC MEANS to calculate means using SAS Version 9.1 (SAS 2002).

Differences between sites or between treatments (monitoring methods) were determined

by one-way analysis of variance (ANOVA) using Fisher's categorical test where

populations were not normally distributed (PROC ANOVA) (SAS 2002). Correlations

among sticky, spot and alsynite monitoring methods were analyzed using a three-way

correlation analysis (Pearson's coefficient) analysis with Dairy A data from wk 15-23

when all three monitoring methods were in use (PROC CORR, SAS 2002). Paired

correlations between spot and sticky cards were performed using Pearson's coefficient









with Dairy B data from weeks 15-23 when both methods were in use (PROC CORR,

SAS 2002). Correlation analysis was conducted for spot card data collected during weeks

that cards were in use at both dairies. Scudder grid and sweep net counts were subjected

to correlation analysis (PROC CORR, SAS 2002). Meteorological data (BP, RH and air

temperature) and data from Scudder grid and net sweeping were collected simultaneously

at Dairy A during weeks 1-9. These data were subjected to correlation analysis (PROC

CORR, SAS 2002).

Enumeration of both aerobic bacteria and of E. coli 0157:H7 included calculations

of the range, mean and median values using spreadsheet functions (Excel 2003). The

number of colony forming units of plates containing 14-300 CFU/plate was multiplied by

the dilution factor, and the count was recorded in scientific notation to one significant

digit. Minimum and maximum values of individual sample types on different days were

not examined statistically.

Prevalence rates of E. coli 157:H7, i.e., the percent of tested samples that tested

positive with at least one positive isolate, were calculated for presumptive identification

on both CHROMAgar and CT-SMAC agar plates using spreadsheet functions (Excel

2003). Similarly, PCR confirmation of prevalence rates of presumptive positive samples

from both direct culture agars was calculated using spreadsheet functions (Excel 2003).

Prevalence rates were calculated by dividing the number of presumptive-positive samples

by the total number of tested samples, and multiplying by 100.

Active fly monitoring methods were examined for relationship trends to

Escherichia coli 0157:H7 prevalence rates on CHROMAgar using spreadsheet functions

(Excel 2003). CHROMAgar prevalence rates were used rather than PCR prevalence rates









because direct culture methods were performed on fresh samples while PCR was

performed on presumptive positive isolates that had been stored for approximately 1.5 yr.

Also, because only positive isolates of samples which were presumptive positive using

direct culture methods were submitted to PCR, the number of samples submitted to PCR

was less than the original number of samples that were examined microbiologically. Data

which suggested the existence of trends were then analyzed for correlations statistically

using SAS with Pearson's coefficient, as appropriate (PROC CORR, SAS 2002).

Results

Fly Monitoring

There were no correlations at Dairy A between alsynite traps and spot cards or

between alsynite traps and sticky cards in either the three-way or paired analyses during

the nine weeks that all three devices were used at Dairy A. Positive correlation

(r=0.67444, p = 0.0463) (r2 = 0.4548) between spot cards and sticky cards was observed

when data from 30 October 2008 to 4 December 2008 were subjected to three-way

analysis. A positive but weaker correlation (r=0.42943, p<0.0001) (r2 = 0.1844) was

observed for paired analysis of card data from 23 September 2008 to 4 December 2008.

During the same nine weeks at Dairy B, there was a positive correlation (r=0.69895,

p=0.0362) between data from alsynite traps and spot cards. Sticky cards were not used at

Dairy B.

There was a correlation of spot card numbers between dairies when all spot cards,

including cards from the milk barn, were used in the analysis (F=16.06; df=1,418;

P<0.001). There was stronger correlation between the spot card counts at different dairies

when Dairy A's milk barn data were excluded from the analysis (F= 55.58; df=1,323;

P<0.0001).









Sweep net fly counts were higher than fly counts on Scudder grids, but there was

no correlation between the two active fly monitoring methods even when data were

logarithmically transformed. Scudder grid counts and sweep net counts were not

influenced by weather conditions (barometric pressure, relative humidity, or

temperature).

Enumeration of Aerobic Bacteria and Escherichia coli 0157:H7

Over the entire study period from 14 June 2008 to 16 September 2008, 35 house

fly, 24 spilled grain and nine manure samples were collected. Of these 68 samples, 14 fly,

17 grain and six manure samples were obtained from Dairy A while 10 fly, seven grain

and three manure samples were obtained from Dairy B. During the same period seven

and four fly samples were collected from Restaurants C and D, respectively. Because

selective media were used for enumeration of both aerobic bacteria and E. coli 0157:H7,

actual counts might be underreported.

Enumeration of aerobic bacteria varied across sample types and dates (Table 3-3).

All sample types contained at least one sample containing 107 aerobic bacteria (Table 3-

3). Mean aerobic plate counts for house flies, grain, and manure samples for the entire

study period were 5.1 x106, 2.0 x 107, and 2.1 x 107 CFU/g, respectively (Table 3-4).

Overall aerobic bacteria counts in grain and manure samples were 10-100-fold greater at

Dairy A than at Dairy B (Table 3-4). House fly carriage of aerobic bacteria was similar in

magnitude for both dairies and Restaurant C (Table 3-4). Flies from Dairy A had many

more bacteria than flies from Dairy B (Table 3-4), which is consistent with higher grain

and manure bacterial loads at Dairy A than at Dairy B. Flies from Restaurant D had 100-

fold fewer bacteria than from all other sites (Table 3-4). However, aerobic bacteria were

enumerated from only one Restaurant D house fly sample, whereas two samples from









each dairy and three samples from Restaurant C were enumerated. Dairy A flies averaged

8.5 x 106 aerobic bacteria and Dairy B flies averaged 1.1 x 106. Manure samples at Dairy

A averaged 3.7 x 107 aerobic bacteria while manure samples from Dairy B averaged 4.6 x

106. At each dairy, aerobic bacteria counts were lower in flies than in substrate media.

Dairy A grains had higher aerobic bacterial loads than manure, but Dairy B grains had

lower bacterial loads than manure.

Enumeration ofE. coli 0157:H7 for 31 May 2008 and 14 June 2008 ranged from

2.7 x 105 to 1.0 x 107 CFU/g E. coli 0157:H7. These data were generated from one house

fly and one manure sample on the former date and one grain and onemanure sample on

the latter date.

Escherichia coli 0157:H7 Prevalence by Direct Culture

A total of 68 samples were collected from all sites, of which 57 samples were

tested using CHROMAgar plates. Collected samples consisted of 35 house fly, 24 grain,

and nine manure samples. Tested samples were comprised of 33 fly, 17 grain, and seven

manure samples. Time constraints prevented testing of the remaining 11 collected

samples. Microbiological processing of the 57 samples produced 197 isolates that were

comprised of 103 fly, 61 grain, and 33 manure isolates.

Of the 57 tested samples, 11 (19.3%) were presumptive positive for E. coli

0157:H7 (Table 3-5) on CHROMAgar. Across the study, E. coli 0157:H7 combined

sample prevalence for both dairies was 17.4% (8/46) with 13.0% (6/46) from Dairy A

and 4.3% (2/46) from Dairy B. Combined prevalence ofE. coli 0157:H7 at the

restaurants was 27.3% (3/11), with 18.2% (2/11) from Restaurant C and 9.1% (1/11)

from Restaurant D. Escherichia coli 0157:H7 was presumptively isolated and identified

from house flies at all four sites and from grain at both dairies using CHROMAgar plates









(Table 3-5). All manure samples tested negative on CHROMAgar. Respective overall

presumptive positive prevalence rates of E. coli 0157:H7 isolated from house flies, grain,

and manure samples were 14.0% (8/57), 5.3% (3/57) and 0% (0/57).

Within each tested sample type, 24.7% (8/33) of the flies, 17.6% (3/17) of the

grain, and 0.0% (0/7) of the manure samples were presumptive positive on

CHROMAgar. Within samples collected only at farms, 18% (5/28) of the samples at

Dairy A and 11% (2/18) of the samples at Dairy B were presumptive positive on

CHROMAgar. There was no relationship between E. coli 0157:H7 presumptive positive

prevalence rates and the Scudder grid fly counts or the sweep net fly counts.

Similarly, samples that were positive on CT-SMAC agar within each tested sample

type were comprised of six fly, five grain, and two manure samples. These 13 samples

tested positive on CT-SMAC agar only, while nine additional samples that tested positive

on CT-SMAC were clones of CHROMAgar positive samples. Within samples collected

only at farms, 6/9 from Dairy A, 3/4 from Dairy B, 2/2 from Restaurant C, and 1/2 from

Restaurant D were presumptive positive on CT-SMAC agar.

Escherichia coli 0157:H7 Prevalence by Polymerase Chain Reaction

As described previously, isolates of all 11 samples that were presumptive positive

on CHROMAgar for the entire study were submitted to PCR (Table 3-5). Additionally,

13 samples that were presumptive positive on CT-SMAC agar plates on 26 August 2008

and 16 September 2008 were submitted to PCR. Thus, 24 total samples were submitted to

PCR. Twelve of 24 (50%) samples were positive using multiplex PCR. The number of

samples positive for both gene fragments was increased to 14/24 (58%) when PCR was

done separately for each primer pair. Samples obtained in wk 6 (5 August 2008) were not

available for PCR assays due to freezer thawing that killed the corresponding isolates.









Specific breakdown of PCR confirmation by sample type of the samples that were

submitted to multiplex PCR analysis revealed that 58% (7/12) of house flies, 56% (5/9)

of grain and 0% (0/3) of manure samples were confirmed as positive. Additional testing

by uniplex PCR increased the confirmation rates to 67% (8/12) for house flies and (1/3)

for manure samples, while prevalence rates for grain were unchanged. Overall prevalence

increased from 50% (12/24) to 58% (14/24).

Nanodrop spectrophotometry analysis resulted in 260:280 ratios that ranged from

1.82 to 1.90. The quantity of DNA template extracted from cultures ranged from 2-9

ng/pl in the four isolates (31 May 2008 and 14 June 2008) that were evaluated using

spectophotometry nanodrop analysis.

The 16S rDNA efficiency of the 46 selected isolates was 89% (41/46 isolates).

Eleven percent of the samples did not contain detectable levels of DNA. This suggests

that up to 11% of the 197 isolates (corresponding to the original 57 samples) that did not

produce amplicons of the expected size might have contained degraded DNA.

Discussion

Sweep net fly counts were generally higher than Scudder grid fly counts. This

supports the findings of Dhillon and Challet (1985), who found that fly counts with

sweep nets were nearly double fly counts on Scudder grids. However, they did not report

an examination of their data for correlation between the two fly monitoring methods.

Some challenges were experienced during this study. In the field, house fly

populations were highly variable throughout the sampling period for all monitoring

methods. This was particularly true for the restaurant garbage dumpsters in town, making

collection of adequate numbers of flies difficult. In many instances, this may have been

caused by insecticide bait or residual insecticide applications, as large numbers of dead









and/or twitching flies accompanied by fly bait pellets were observed at all sites on several

occasions. Therefore, the microbiological protocol used for processing fly samples in this

research had to be modified to accommodate collection of smaller numbers of flies than

originally planned. Decreased numbers of flies in pools did not appear to have a

detrimental effect upon isolation ofE. coli 0157:H7, as positive fly samples were

obtained continuously throughout the test period.

During warmer months (summer and fall), sweep-netted flies placed in the cooler

could not be sufficiently knocked down to allow for field identification and sorting of

individual flies into microcentrifuge tubes. This was likely caused by the high ambient air

temperatures, which rapidly melted the ice in the cooler. As a result of these constraints,

collection of flies was conducted differently during cold and hot months. Modification of

the fly collection protocol may have influenced the results of this, because flies collected

in warmer months remained relatively active for 2-6 h. This may have resulted in

artificial or increased transfer of pathogenic microorganisms among flies held in cups that

could have contributed to the high prevalence rates that I obtained from house fly

samples. The use of dry ice in coolers during hot weather would lessen this effect, but dry

ice production of CO2 might adversely impact aerobic growth, so that another method of

implementing steady cool temperatures would be useful.

House fly carriage of aerobic bacteria was variable between all sites with

restaurants providing counts at both extremes. At each dairy, aerobic bacteria counts were

highest in manure and lowest in flies. This supports the possibility, as reported by (Vold

et al. 2000) that high background counts of competing microorganisms decreases

detection of E. coli 0157:H7. In particular, because isolation of E. coli 0157:H7 from









dairy cattle manure was so low in comparison to that from house flies and spilled grain,

the relationship of aerobic bacteria with E. coli 0157:H7 needs to be examined further,

within the various media that were tested in this experiment. Additionally, the role of

house flies as a potential reservoir, not just as pathogen vectors, needs to be studied more.

Enumeration of E. coli 0157:H7 ranged from 2.7 x 105to 2.4 x 106 for the two

samples obtained on 31 May 2008 (one house fly and one manure) and the two samples

obtained on 14 June 2008 (one grain and one manure). On 31 May 2008, the house fly

sample from Dairy A contained 2.4 x 106 CFU/g. These data are in agreement with E.

coli 0157:H7 and aerobic bacteria counts reported previously from house flies (Alam and

Zurek 2004, Sanderson et al. 2005), cattle feed (Ahmad et al. 2007), and cattle feces

(Brichta-Harhay et al. 2007).

The main purpose of this study was to determine prevalence rates of E. coli

0157:H7 at two dairy farms and in two restaurant garbage dumpsters in a nearby town.

Escherichia coli 0157:H7 was isolated from house flies at all four locations, and from

grain at both dairies. Although no E. coli 0157:H7 was isolated using direct culture

methods from manure, only seven manure samples were tested, whereas 17 grain and 33

house fly samples were tested. Lahti et al. (2003) and Omisakin et al. (2003) observed

that detection was directly linked to numbers of samples processed. If target pathogen

numbers are low, then detection can be difficult without increased numbers of samples

(Brichta-Harhay et al. 2007). Thus, it is possible that E. coli 0157:H7 might have been

detected in manure if more samples had been processed. Conversely, recovery ofE. coli

0157:H7 from manure can be inconsistent due to high densities of competing

background microorganisms (Pao et al. 2005), so that increasing the number of samples









might have had no impact on prevalence rates. Zero or very low recovery rates are not

uncommon in cattle feces. Hancock et al. (1994) recovered E. coli 0157:H7 from only

0.28% (10 of 3,570) of dairy cattle feces samples and Galland et al. (2001) isolated E.

coli 0157:H7 from only 0.26% (45/17,050) of cattle fecal pats. Overall prevalence of E.

coli 0157:H7 at both dairies combined ranged from 3.6 10.7%, while prevalence at the

restaurants combined ranged from 1.8 7.1 %. These data are in agreement with isolation

rates obtained at dairies in some studies (Heuvelink et al. 1998, Bonardi et al. 2001,

Smith et al. 2005, Oporto et al. 2008), although not as high as reported by others

(Sanderson et al. 2006).

The results obtained in this study with respective overall prevalence rates for

house flies, grain, and manure samples at 14.0% (8/57 samples), 5.3% (3/57), and 0.0%

(0/57) support the findings of previous authors (Lahti et al. 2003, Pao et al. 2003, Pearce

et al. 2004, Brichta-Harhay et al. 2007) that increased sampling may be an important

factor in pathogen detection. Direct culture isolation ofE. coli 0157:H7 from house flies

was approximately 2.6 times greater than from grain. This suggests that detection ofE.

coli 0157:H7 on dairies might be more accurately determined by testing house flies

instead of grain or manure samples, regardless of which isolation method is utilized. In

addition to providing higher prevalence ofE. coli 0157:H7, house flies can carry and

excrete this pathogen for up to 4 d (Sasaki et al. 2000). Additionally, house flies can

disperse from dairies to restaurants and other sites in town up to 3.0 km distant (Chapter

2).

Recovery of 24.6% of E. coli 0157:H7 from house flies is a much higher

recovery rate than those reported by Agui et al. (2001) (7.2%) and Keen et al. (2006)









(5.2%), but less than that that reported by Fotedar et al. (1992) (31.2-33.8%). There were

substantial differences in the culture and isolation methods used by Augie et al. (2001)

and Keen et al. (2006) versus those methods used by Fotedar et al. (1992) and myself.

Agui et al. (2001) and Keen et al. (2006) used enrichment broths that were selective for

E. coli. They added multiple antibiotics to the selective enrichment broths and also to the

CHROMAgar plates. In contrast, both Foetedar et al. (2006) and I used a non-selective

enrichment broth, to which only one antibiotic, instead of multiple antibiotics, was added.

Also, I did not supplement the CHROMAgar plates with antibiotics. This suggests that

antibiotics might be inhibitory for isolation ofE. coli 0157:H7 from house flies. Thus,

testing house fly samples instead of grain and manure at dairies might provide a cost-

savings to researchers in addition to higher prevalence rate data, because expenses

associated with purchasing, preparing, storing, and disposing of antibiotics can be greatly

reduced. Elimination of antibiotics from broths and agars would extend the shelf life of

these products, further increasing cost savings due to reduced labor expenses.

Although CHROMAgar 0157 is very selective and specific for E. coli 0157,

violet-colored colonies presumptive for E. coli 0157:H7 tended to grow slowly in this

experiment. Slow growth of presumptive colonies on CHROMAgar plates was

discovered when one batch of negative plates remained on the bench for 24 h after

removed from the incubator. Previously, CHROMAgar plates that had not shown positive

presumptive growth were discarded after the initial 24 h incubation. After this discovery,

CHROMAgar plates were retained and reexamined at both 24 and 48 h after removal

from incubator. Prevalence of E. coli 0157:H7 on CHROMAgar plates may have been

under-reported in this study because most samples were subjected to 6-18 h incubation.









The final batch of CHROMAgar that was used for isolation on 16 September

2008 was excessively diluted so that the agar was difficult to streak. Colonies did not

remain on top, but sank into the media. Because the typical morphology and color was

not discernible, all CHROMAgar plates on 16 September 2008 were recorded as

negative. However, up to five isolates were transferred onto CT-SMAC plates by digging

a loop into "submerged" CHROMAgar colonies and streaking as usual. Seventeen of 75

isolates transferred from the "negative" CHROMAgar plates to CT-SMAC plates tested

positive on CT-SMAC. This clearly suggests that the colonies may have been positive on

the CHROMAgar media, despite the improperly-made media that obscured normal

reading of plates. The 17 positive isolates were distributed from the 10 different samples,

so that all samples tested positive in CT-SMAC with at least one positive isolate.

However, all samples for this date were recorded as negative on CHROMAgar for this

study (Tables 3-5).

It is also possible that competing organisms grew on the CHROMAgar in

sufficient quantities to outcompete E. coli 0157:H7. Although designed for selective

growth ofE. coli 0157:H7, closely related coliforms including Proteus spp. can grow.

False positives are also a possibility on CHROMAgar that is not supplemented with

potassium tellurite, because non-E. coli 0157:H7 bacteria such as Salmonella spp. have

the same colony color on this media (CHROMAgar protocol, Invitrogen, CA). More

closely related results between CHROMAgar and CT-SMAC plates might have been

obtained if the CHROMAgar had been supplemented with potassium tellurite. However,

this study has higher prevalence rates than studies that did supplement CHROMAgar









with antibiotics, so that the impact of antibiotics upon isolation ofE. coli 0157:H7 has

yet to be fully understood.

One of the original goals of this study was to determine if there were correlations

between fly population densities and E. coli 0157:H7 prevalence using direct culture

methods at Dairy A. However, due to changes in farm management practices on Dairy A,

this became a minor research topic. Specifically, the location for feeding the cattle was

changed approximately halfway through the study from the milk barn to the feed barn.

Therefore, only active fly monitoring methods could be examined for potential

correlations to E. coli 0157:H7 prevalence, because Scudder grids and sweep netting

were consistently conducted at Dairy A's feed barn, the microbiological specimen

collection site. In contrast to active fly monitoring methods, it was not possible to look

for correlations of passive fly monitoring methods to E. coli 0157:H7 prevalence because

spot cards and sticky cards were not in use during the entire microbiological sampling

period.

Another goal of this study was to use PCR assays to confirm direct-culture

presumptive isolation and identification of E. coli 0157:H7 colonies conducted in this

project. In this study, only 58% (14/24) of all samples that were presumptive-positive

using direct-culture media were confirmed by PCR analysis using both multiplex and

uniplex assays, despite using multiple isolates for samples. The 14 samples that were

confirmed as positive by PCR comprise only 25% of the initial 57 samples that were

cultured. Confirmation by PCR of only 58% of the samples suggests that the remaining

42% were either false positives on culture plates or that the fresh cultures lacked adequate









quantities of DNA for successful PCR amplification following the extended time in the

freezer.

It is interesting that 33% (1/3) of manure tested positive by uniplex PCR, because

all three manure samples that were tested by PCR were obtained from isolates that

appeared negative using CHROMAgar media. These results appear to confirm either an

increased sensitivity of PCR over direct culture (Fratamico et al. 2005) or detection of

sorbitol-fermenting strains (Cebula et al. 1995). As with direct culture methods, PCR

testing of house flies in this study provided the best method for detecting E. coli 0157:H7

on dairies, and provided more information than grain or manure samples did about the

presence ofE. coli 0157:H7 on the dairies. Therefore, house flies should be an important

part of any sampling program at dairies when looking for this pathogen.

Confirmation by PCR might have been reduced in this study because purified

isolates were stored long-term at -20 OC, so that thawing and refreezing of cultures might

have contributed to mortality (Mennigmann 1979), gene loss (Acha et al. 2005), and/or

contamination by laboratory microorganisms (V. Mai, personal communication). In this

study, an 8.3% mortality rate was observed over 1.5 yr, which is similar to 7% mortality

over nine mo reported by Doyle and Schoeni (1984). Mortality of individual isolates

likely impacted the PCR confirmation rate, and underscores the importance of obtaining

multiple isolates from each presumptive positive colony. Survival of cultures in -20 C

storage was recently shown to be increased by restricting the exponential growth phase to

approximately 3 h or increasing the nutrient availability prior to storage (Sezonov et al.

2007). Cultures in this study were provided with minimal nutrients and were incubated









for 24+2 h, so that they might have been dead or stressed and nonviable prior to their

placement in the freezer (Sezonov et al. 2007).

Samples positive forfliCH7 gene segments could include E. coli of serotypes other

than 0157 (Cebula et al. 1995, Mead and Griffin 1998, Szalanski et al. 2004). Because

the combination offliCH7 and rfbEo157 is unique to E. coli 0157:H7 (Bilge et al. 1996), a

multiplex PCR assay including these two fragments was desired. I obtained only one

band in several isolates using multiplex PCR. For samples that showed only one band, I

subsequently performed uniplex PCR for each gene fragment separately. In this manner, I

detected the presence of genes in isolates that had previously not produced a band that

was intense enough to be visualized on the multiplex agarose gels. The lack of gene

fragment amplification in the multiplex PCR could have been due to the depletion of

nucleotides by the other primer, particularly if the primer pair for one gene was working

more efficiently than the other pair.

It was difficult to establish a suitable annealing temperature that worked well for

both gene fragments. Initial attempts at multiplex PCR were conducted without success

using the protocol with an annealing temperature of 48 C published by Szalanski et al.

(2004). As soon as the annealing temperature was increased to > 55 C, desired gene

fragments were successfully amplified during PCR assays.

Bacterial 16S rDNA was detected by PCR for 41 of 46 (89%) samples verifying

that the gene fragments obtained in the multiplex and uniplex PCR assays were due to

bacterial DNA, because rDNA, located on ribosomal genes, contains nucleotide

sequences that are highly conserved in all bacteria species. However, five samples (11%)

tested negative for 16S rDNA. This can occur if the samples are contaminated with









proteins or if DNA concentrations are low or sample inhibitors were not removed during

DNA purification. Because DNA absorbs ultraviolet (UV) light at 260 nm and protein

absorbs UV light at 280 nm, pure DNA samples will exhibit a 260:280 ratio of 1.8 2.0

(Altshuler 2006), while protein-contaminated samples exhibit 260:280 ratios lower than

1.8 (Altshuler 2006). The 260:280 ratios for the four samples processed with this

technique ranged from 1.82 to 1.90, which indicated that this representative selection of

samples was relatively free from protein contamination.

Although relatively free from protein contamination, the concentrations of DNA

after extraction ranged from 2- to 9-ng/pl, which is low. PCR testing of fecal specimens

can be very difficult, because many PCR-inhibiting substances are extracted from

samples along with the target DNA (Holland et al. 2000). Inhibiting factors commonly

found in fecal samples include bile salts, heme, bilirubins, and complex carbohydrates

(Holland et al. 2000). Because my samples were cultured on sensitive media

supplemented with antibiotics, the impact of PCR-inhibiting factors in the PCR assays

should have been minimal. All manure samples that were submitted to PCR analysis were

culture-negative, indicating that perhaps fecal background did inhibit direct-culture

detection. In the current study, PCR was able to detect one of those samples as positive,

due to the combined presence of both target gene fragments. This illustrates the

sensitivity of the PCR over direct-culture, and emphasizes the importance of performing

both techniques. Results of PCR analysis of samples in this study cannot be directly

contrasted with direct-culture analysis because the two methods were not used

simultaneously on fresh samples.









To my knowledge, this is the first study where attempts were made to correlate

house fly bacteria loads to house fly population dynamics on dairies. Enumeration of E.

coli 0157:H7 in this study was insufficient to fully explore this relationship, but future

studies of this suspected correlation could provide a valuable public health and veterinary

health tool. If a correlation exists between house fly populations and pathogen

prevalence, then the epidemiological role of the house fly in a disease transmission cycle

would be better understood, and possibly better quantified in terms of economic and

health costs. This would emphasize the importance of having a strong IPM program for

flies. Additionally, detection ofE. coli 0157:H7 at dairies and other livestock-rearing

facilities will continue to be important, particularly as human urban and suburban

residential areas continue to expand. Flies provided the most reliable source ofE. coli

0157:H7 from dairies, and future research should explore direct detection ofE. coli

0157:H7 from house flies to better understand the role that the house fly might have in

dissemination of this pathogen. Finally, the interactions of aerobic bacteria and E. coli

0157:H7 within different media on a dairy farm need to be examined more, to

understand why E. coli 0157:H7 is more prevalent in house flies than in dairy cattle

manure.

































Figure 3-1. Scudder grid (45 x 45 cm) used to assess house fly populations on dairy
farms. Scudder grids were placed on top of spilled grains on the ground or on
top of feed troughs. Fly counts consisted of the numbers of house flies that
were resting on the grid five seconds after placement. Up to 10 Scudder grid
counts were performed at each site. Scudder grids were placed in the same
location at approximately the same time of day by the same operator to
minimize variability.

























4 mall
1* 9 7Tr16


Figure 3-2. Spot cards at Dairy A's milk barn and at Dairy B were inserted into metal
frames which were nailed to horizontal wooden support beams 2-3 m above
the ground. Cards were held in place by a small metal binder clip placed at the
top edge of the metal frame.











































Figure 3-3. Spot and sticky cards. Spot card clipped in place by binder clips at Dairy A's
feed barn at a height of 2 m above the ground. Sticky card clipped in place
with two paper clips 1 cm below the spot card.










3--


0 A


B











C

Figure 3-4. Sampling methods for each type of collected sample. A) 100 g of grain or
manure was placed in specimen cups. B) Two swabs were pre-moistened with
Buffered Peptone Water (BPW), exposed to grain or manure and placed in 9
ml BPW in a centrifuge tube. C) House flies that were captured using sweep
nets were transferred into specimen cups.









Table 3-1. Dates and house fly monitoring methods used at two Florida dairies.
Date Dairy A1 Dairy B
7 June 2008 to 23 September 2008 Sweep nets, (FB) Sweep nets
Scudder grids, (FB) Scudder grids
Spot cards, (MB) Spot cards
23 September 2008 to 4 December 2008 Spot cards, (FB) Spot cards
Alsynite traps Alsynite traps
30 October 2008 to 4 December 2008 Sticky cards, (FB) N/A
14 May 2009 to 25 June 2009 Alsynite traps Alsynite traps

1Monitoring on Dairy A was conducted at both the milk barn (MB) and the feed barn
(FB). Due to discontinued use of the milk barn by the producer, the milk barn was
eliminated from this study after 23 September 2008.











Table 3-2. Primer nucleotide sequences used to amplify target genes in PCR assay.
Expected PCR Target
Primer1 Sequence (5'-3') amplicon (bp) gene


Assay 1


(multiDlex: 0157 somatic and H7 flagellar antigens)


fliCH7F GCG CTG TCG AGT TCT ATC GAG C
fliCH7R CAA CGG TGA CTT TAT CGC CCA TTC C
rfbEos57-F CGG ACA TCC ATG TGA TAT GG
rfbEoi57-R TTG CCT ATG TAC AGC TAA TCC
Assay 2 (uniplex: 0157 somatic antigen)
rfbEoi57-F CGG ACA TCC ATG TGA TAT GG
rfbEo157-R TTG CCT ATG TAC AGC TAA TCC
Assay 3 (uniplex: H7 flagellar antigen)
fliCH7F GCG CTG TCG AGT TCT ATC GAG C
fliCH7R CAA CGG TGA CTT TAT CGC CCA TTC C

1 F, forward primer; R, reverse primer.
2 Reference: (1) (Gannon et al. 1997); (2) (Paton and Paton 1998)


625

259


259


625


flic

rfb


rfb


flic


Specificity

H7 flagellar gene

nt 393-651 ofr/Jbo57:H7


nt 393-651 ofr bol57:H7


H7 flagellar gene


Citation2









Table 3-3. Enumeration of aerobic bacteria (CFU/g) using Petrifilm Aerobic Plate Count
plates inoculated with 1 ml of the unenriched sample.
Aerobic Plate Count
Date Site, Source Range (CFU/g) Mean Median
(CFU/g) (CFU/g)
5/31/08 A, Manurea, 2.5 x 106 2.5 x 106 2.5 x 106
6/14/08 A, Grain 9.8 x 107 9.8 x 107 9.8 x 107
A, Manure 2.8 x 10 4.6 x 107 3.7 x 107 3.7 x 107
B, Grain 4.7 x 106 4.7 x 106 4.7 x 106
B, Manure 2.5 x 106 -6.7 x 106 4.6 x106 4.6 x106
C, House fly 1.3 x 107 3.0 x 107 2.2 x 107 2.2 x 107
6/23/08 A, Grain 1.6 x 10 2.8 x 106 1.5 x 106 1.5 x 106
A, House fly 1.4 x 10 2.0 x 107 1.7 x 107 1.7 x 107
B, Grain 1.8 x 105 3.0 x 105 2.4 x 105 2.4 x 105
B, House fly 2.2 x 10 4.0 x 106 2.1 x 106 2.1 x 106
C, House fly 1.3 x 103 1.3 x 103 1.3 x 103
8/26/08 A, Grain2 4.8 x106 8.2 x 107 4.3 x 107 4.3 x 10
A, House flyb 3.4 x 104 -1.5 x 105 9.2 x 104 9.2 x 104
B, Grain 2.9 x 106 3.5 x 106 3.2 x 106 3.2 x 106
B, House fly 1.1 x 105- 1.4 x 105 1.3 x05 1.3 x105
C, House fly 1.5 x 104 1.5 x 104 1.5 x 104
D, House fly 2.1 x104 2.1 x 104 2.1 x 104

Samples were collected from two dairies and dumpsters at two restaurants in north
central Florida. A=Dairy A, B=Dairy B, C=Restaurant C and D=Restaurant D.
a Samples were enumerated for aerobic plate counts, but not tested for E. coli 0157:H7.
b Two samples tested. Remaining data are CFU/g per plate for single samples.











Table 3-4. Mean enumeration, by site and by sample type, of aerobic bacteria (CFU/g) using Petrifilm Aerobic Plate Count plates
inoculated with 1 ml of unenriched samples enumerated from collections between 31 May 2008 to 26 August 2008 on
dairies and at restaurants in north central Florida.
Mean Aerobic Plate Count (CFU/g)
Substrate Dairy A Dairy B Dairiesa Restaurant C Restaurant D Restaurantsa All flies
Grain 3.8 X 107 2.3 X 106 2.5 x 107 NA NA NA NA
Manure 3.7 X 107 4.6 X 106 2.1 x 107 NA NA NA NA
Housefly 8.5 X 106 1.1 X106 4.8 x 106 1.1x 10 2.1 x 104 5.4 x 106 5.1 x 106

a Means for dairies and restaurants were calculated by adding the means of Dairy A and Dairy B, or of Restaurant C and Restaurant
D, respectively, and dividing the result by two. Means for all flies were calculated by dividing the summed means of all four sites
and dividing by four. Thus, means for dairies, restaurants and all flies were simple, not weighted.
NA, Not applicable.









Table 3-5. Prevalence (%) and number of E. coli 0157:H7 CHROMAgar-positive samples and number of CT-SMAC and PCR-
positive samples relative to the number of samples tested after collection from two dairies and two restaurant dumpsters in
north central Florida.


CHROMAgar CT-SMAC PCR

Sample No. Positive
Date Site Type No. Collected No. Tested (%) No. Tested No. Positive No. Tested No. Positive


6/14/2008


A

B


C
6/23/2008 A

B

C
7/20/2008 A
B
C
7/28/2008 A
B
C
D
8/5/2008 A


Grain
Manure
Grain
Manure
Fly
Fly
Grain
Fly
Grain


2
2
2
2


1
1
1
1


Fly
Fly
Fly
Fly
Grain


Manure
B Fly
Grain


1(100)
0 (0)
0 (0)
0 (100)
0 (0)
0 (0)
0 (0)
0 (0)
0 (0)
0 (0)
1(50)
0(0)
1(100)
0 (0)
0 (0)
0 (0)
0 (0)
1 (33)
0 (0)
0 (0)
0 (0)
0 (0)


1
1
1
1
1
NA
NA
NA
NA
NA
1
0
1
NA
NA
NA
NA
NA
NA
NA
NA
NA


1
NA
NA
NA
NA
NA
1
NA
1
NA
NA
NA
NA
NA
NA
NA
NA
NA


NA
NA
NA
NA
NA




NA
NA
NA
NA
NA
NA
NA
NA
NA











CHROMAgar CT-SMAC PCR

Sample No. Positive
Date Site Type No. Collected No. Tested (%) No. Tested No. Positive No. Tested No. Positive
Manure 1 1 0(0) NA NA NA NA
C Fly 1 1 1(100) NA NA NA NA
D Fly 1 1 0(0) NA NA NA NA
8/26/2008 A Fly 5 5 2(40) 5 2 2 1
Grain 6 6 1(17) 6 2 5 3
B Fly 1 1 1(100) 1 0 1 1
Grain 1 1 1(100) 1 1 1 1
C Fly 1 1 0(0) 1 1 1 1
D Fly 1 1 1(100) 1 1 1 1
9/16/2008 A Fly 1 1 0 (0) 1 1 1 1
Grain 2 2 0 (0) 2 2 1 1
Manure 2 2 0 (0) 2 2 1 1
B Fly 1 1 0(0) 1 1 1 1
Grain 2 2 0 (0) 2 2 2 0
C Fly 1 1 0(0) 1 1 1 1
D Fly 1 1 0(0) 1 1 1 1
TOTAL 68 57 11(19.2) 32 22 24 14

Sites: A, Dairy A; B, Dairy B; C, dumpster at Restaurant C; D, dumpster at Restaurant D. Spilled rain and manure samples
consisted of either 25 g substrate or swabs. Fly samples consisted of pools of up to 25 house flies. Distance (km) from release
site (Chapter 2) is also provided. CT-SMAC, sorbitol MacConkey agar supplemented with cefixime and potassium tellurite.
a Percent positive was calculated by dividing the number of positive samples by the number of samples tested in the laboratory.









CHAPTER 4
OVERALL CONCLUSIONS

Conclusions

The overall goal of this research was to determine the role of house flies, Musca

domestic L., in the transmission ofEscherichia coli 0157:H7 at the rural-urban interface using

two dairies and a small town. This was accomplished through three sampling parameters: by

examining house fly dispersal, house fly population patterns, and Escherichia coli 0157:H7

prevalence. Specifically, studies were conducted to determine if house flies disperse from dairies

into town, what their populations on the farms were at the time of the dispersal studies, and

characterizing E. coli 0157:H7 prevalence from house flies at both the dairies and in town using

CHROMAgar and sorbitol MacConkey agar (CT-SMAC) direct culture methods. Additionally, a

rapid, multiplex polymerase chain reaction (PCR) method was developed to detect the rfbEH7 and

fliCo157 virulence factor genes of E. coli 0157:H7 isolated from house flies, grain and manure.

Escherichia coli 0157:H7 prevalence data was used in conjunction with house fly dispersal data

from the dairies to the town to estimate the public health risk that house flies may present in

regards to transmission of E. coli 0157:H7 from dairies into towns located in north-central

Florida.

Background

Previous research documented E. coli 0157:H7 isolation from house flies on dairies

(Zhao et al. 1995) and at restaurants (Butler et al. 2010). Laboratory studies have shown that E.

coli 0157:H7 survives and is excreted in a viable, infectious state for up to 4 d after it is ingested

by house flies (Sasaki et al. 2000), while Petridis et al. (2006) and Macovei et al. (2008)

demonstrated replication ofE. coli 0157:H7 in house flies. Field studies have shown that house

flies can disperse up to 4.8 km (Quarterman et al. 1954, Shura-Bura et al. 1962) and that flies are









capable of flying at rates of 1 km/h (Shura-Bura et al. 1962). However, no studies have been

conducted to investigate the synergism of house fly dispersal and E. coli 0157:H7 survival in the

natural environment.

Successful isolation of this pathogen from flies located both at the dairies and in town

followed by genetic analysis of both the flies and the bacteria would provide more information

about origin of bacterial strains isolated from flies. Isolation by direct culture reveals important

information about the biochemical characteristics of bacteria, but isolation by DNA-based

methods such as PCR permit serotyping and phylogenetic analysis. In addition, PCR is more

sensitive and specific than direct culture techniques. Both direct culture and PCR methods have

been used to isolate E. coli 0157:H7 from house flies, grain, and manure samples on dairies.

Pulsed-field gel electrophoresis is the preferred standard for DNA-based identification methods,

but it is more time-consuming and labor-intensive than PCR.

Conclusions

I hypothesize that the house fly provides a mobile element of disease-causing pathogen

transmission from dairy cattle to humans, particularly in locations where the rural-urban

interface is within the typical flight range of house flies. Furthermore, I hypothesize that the role

of house fly pathogen transmission can be better understood by genetic serotyping of both house

fly and bacteria and determining if the same strains exist in both the rural and urban locations

where fly transmission is suspected. I hypothesize that PCR can be used to quickly and

efficiently confirm E. coli 0157:H7 isolation from house flies, grain and manure.

This study provided evidence of house fly dispersal from a dairy into town across the

rural-urban interface, using roads as landscape corridors, and by direct flight overland.

Furthermore, dispersal from a dairy to a restaurant was observed. This study demonstrated the









usefulness of a newly designed multiplex PCR assay for confirmation of direct culture

presumptive-positive E. coli 0157:H7 samples.

Future Research

There are additional studies that need to be conducted to build on the findings of this

research in order to better understand, and perhaps eventually quantify, the role of house flies in

pathogen transmission across the rural-urban interface from dairies into town. In particular, more

efficient DNA-based isolation methods are needed.

In-depth understanding of house fly bacterial loads has been limited by the need to use

fresh samples containing viable bacteria for direct culture isolation. Isolation of bacteria by direct

culture permits long-term storage of viable isolates so that DNA-based analysis can be conducted

at future dates. Therefore, direct culture methods are valuable and should be used. However,

PCR can amplify both viable and non-viable bacteria, so that storage of flies might avoid the

need to culture bacteria samples. This could reduce expenses and labor, as well as increase

specificity and selectivity.

Future research should be conducted to determine if PCR can rapidly and efficiently

isolate E. coli 0157:H7 directly from house fly samples that have been in long-term storage at -

20 C. The relationship between house flies and E. coli 0157:H7 and other enteric bacteria

should be examined for specific geographical regions by using PCR on flies that have been in

long-term storage. Use of flies that have been in long-term storage permits more extensive

sampling during seasons of dominant fly activity because PCR assays can be conducted outside

of the fly season during periods of time when researchers have greater available labor.

Summary

In summary, this research has led to a better understanding of the house fly role in

transmission of E. coli 0157:H7 from dairies to urban areas. This research has resulted in the









development of a multiplex PCR assay for identifying E. coli 0157:H7 from house flies. Results

from this dissertation demonstrate that house flies provide valuable information regarding a

mobile element for pathogen transmission that is lacking in grain and manure samples. House

flies provide a readily-available, high-return sampling option and should be incorporated in the

design of a pathogen monitoring programs on dairies.









LIST OF REFERENCES


Abe, K., S. Yamamoto, and K. Shinagawa. 2002. Economic impact of an Escherichia coli
0157:H7 outbreak in Japan. J. Food Prot. 65: 66-72.

AchA, S. J., I. Kiihn, G. Mbazima, P. Colque-Navarro, and R. Mollby. 2005. Changes of
viability and composition of the Escherichia coli flora in faecal samples during long time
storage. J. Microbiol. Meth. 63: 229-238.

Agui, N. 2001. Flies carrying enterohemorrhagic Escherichia coli (EHEC) 0157 in Japan: a
nationwide survey. Med. Entomol. Zool. 52: 97-103.

Ahmad, A., T. G. Nagaraja, and L. Zurek. 2007. Transmission of Escherichia coli 0157:H7
to cattle by house flies. Prev. Vet. Med. 80: 74-81.

Alam, M. J. and L. Zurek. 2004. Association ofEscherichia coli 0157:H7 with houseflies on a
cattle farm. Appl. Environ. Microbiol. 70: 7578-7580.

Alam, M. J. and L. Zurek. 2006. Seasonal prevalence of Escherichia coli 0157:H7 in beef
cattle feces. J. Food Prot. 69: 3018-3020.

Ananth, G. P., D. C. Bronson, and J. K. Brown. 1992. Generation of airborne fly body
particles by four electrocution fly traps and an electronic fly trap. Inter. J. Environ. Health
Res. 2: 106-113.

Anderson, G. S. and B. J. Danielson. 1997. The effects of landscape composition and
physiognomy on metapopulation size: the role of corridors. Landscape Ecol. 12: 261-271.

Anderson, J. and J. Poorbaugh. 1964. Biological control possibility for house flies. Cal. Agr.
18: 2-4.

Angelo, M. J. and F. Slansky. 1984. Body building by insects: trade-offs in resource allocation
with particular reference to migratory species Fla. Entomol. 67: 22-41.

ANON. 1940. Summer diarrhea and horse-drawn vehicles. Am. J. Pub. Health 30: 825-826.

Archer, D. L. and F. E. Young. 1988. Contemporary issues: Diseases with a food vector. Clin.
Microbiol. Rev. 1: 377-398.

Asakura, H., S. Makino, T. Shirahata, T. Tsukamoto, H. Kurazono, T. Ikeda, and K.
Takeshi. 1998. Detection and genetical characterization of Shiga toxin-producing
Escherichia coli from wild deer. Microbiol. Immunol. 42: 815-822.

Avancini Rita MP and A. R. Silveira Gerson. 2000. Age structure and abundance in
populations of muscoid flies from a poultry facility in Southeast Brazil. Mem. Inst.
Oswaldo Cruz.95: 259-264. (http://www.scielo.br/scielo.php?script=sci_arttext&pid
=S0074-02762000000200022&lng=en).









Avery, L. M., A. P. Williams, K. Killham, and D. L. Jones. 2008. Survival of Escherichia
coli, 0157:H7 in waters from lakes, rivers, puddles and animal-drinking troughs. Sci.
Total Environ. 389: 378-385.

Axtell, R. C. 1970. Integrated fly-control program for caged-poultry houses. J. Econ. Entomol.
63: 400-405.

Bailey, D. L. 1970. Forced air for separating pupae of house flies from rearing medium. J. Econ.
Entomol. 63: 331-333.

Baldwin, T. and E. H. Bryant. 1981. Effect of size upon mating performance within geographic
strains of the housefly, Musca domestic L. Evol. 35: 1134-1141.

Barkocy-Gallagher, G. A., K. K. Edwards, X. Nou, J. M. Bosilevac, T. M. Arthur, S. D.
Shackelford, and M. Koohmaraie. 2005. Methods for recovering Escherichia coli
0157:H7 from cattle fecal, hide, and carcass samples: sensitivity and improvements. J.
Food Prot. 68: 2264-2268.

Barnard, D. R. 2003. Control of fly-borne diseases. Pestic. Outlook 14: 222-228.

Bell, B. P., M. Goldoft, P. M. Griffin, M. A. Davis, D. C. Gordon, P. I. Tarr, C. A.
Bartleson, J. H. Lewis, T. J. Barrett, J. G. Wells, et al. 1994. A multistate outbreak of
Escherichia coli 0157:H7-associated bloody diarrhea and hemolytic uremic syndrome
from hamburgers: the Washington experience. J. Am. Med. Assoc. 272: 1349-1353.

Berg, H. C. 2004. E. coli in motion (biological and medical physics: biomedical engineering).
AIP Press, Springer-Verlag, New York, NY.

Berry, I. L., P. J. Scholl, and J. I. Shugart. 1981. A mark and recapture procedure for
estimating population sizes of stable flies. Environ. Entomol. 10: 88-93.

Bertani, G. 1951. Studies on lysogenesis. I. The mode of phage liberation by lysogenic
Escherichia coli. J. Bacteriol. 62: 293-300.

Bertani, G. 2004. Lysogeny at mid-twentieth century: P1, P2, and other experimental systems. J.
Bacteriol. 186: 595-600.

Bettelheim, K. A. 1991. The genus Escherichia, pp. 2696-2736. In A. Balows, H. G. Truper, M.
Dworkin, W. Harder, and K.-H. Schleifer [eds.], The prokaryotes: a handbook on the
biology of bacteria: ecophysiology, isolation, identification, applications. Springer-
Verlag, New York, NY.

Beutin, L., G. Krause, S. Zimmermann, S. Kaulfuss, and K. Gleier. 2004. Characterization of
Shiga toxin-producing Escherichia coli strains isolated from human patients in Germany
over a 3-year period. J. Clin. Microbiol. 42: 1099-1108.









Beutin, L., A. Miko, G. Krause, K. Pries, S. Haby, K. Steege, and N. Albrecht. 2007.
Identification of human-pathogenic strains of Shiga toxin-producing Escherichia coli
from food by a combination of serotyping and molecular typing of Shiga toxin genes.
Appl. Environ. Microbiol. 73: 4769-4775.

Bilge, S. S., J. C. Vary, S. F. Dowell, and P. I. Tarr. 1996. Role of the Escherichia coli
0157:H7 0 side chain in adherence and analysis of an rfb locus. Infect. Immun. 64:
4795-4801.

Birkemoe, T. A. Soleng, and A. Aak. 2008. Biological control of Musca domestic and
Stomoxys calcitrans by mass releases of the parasitoid Spalangia cameroni on two
Norwegian pig farms. Biocontrol 54: 425-436.

Bishopp, F. C. and E. W. Laake. 1921. The dispersion of flies by flight. J. Agric. Res. 21: 729-
766.

Black IV, W. C. and E. S. Krafsur. 1986. Geographic variation in house fly size adaptation
or larval crowding? Evol. 40: 204-206.

Black IV, W. C. and E. S. Krafsur. 2008. Fecundity and size in the housefly: investigations of
some environmental sources and genetic correlates of variation. Med. Vet. Entomol. 1:
369-382.

[BLAST] Basic Local Alignment Search Tool. 2009. Basic local alignment search tool.
(http://blast.ncbi.nlm.nih.gov/ Blast.cgi).

Bodnaryk, R. P. and P. E. Morrison. 1966. The relationship between nutrition, haemolymph
proteins, and ovarian development in Musca domestic. J. Ins. Physiol. 12: 963-976.

Bolton, D.J., Byrne, C.M., Sheridan, J.J., McDowell, D.A. and Blair, I.S. 1999. The survival
characteristics of a non-toxigenic strain of Escherichia coli 0157:H7. J. Appl. Microbiol.
86:407-411.

Borror, D. J., C. A. Triplehorn, and N. F. Johnson. 1989. An introduction to the study of
insects, 6th ed. Saunders College Publishers, Orlando, FL.

Bosik, J. J. 1997. Common names of insects and related organisms. Entomological Society of
America, College Park, MD.

Boulesteix, G., P. Le Dantec, B. Chevalier, M. Dieng, B. Niang, and B. Diatta. 2005. Role of
Musca domestic in the transmission of multiresistant bacteria in the centres of intensive
care setting in sub-Saharan Africa. Ann. Fr. Anesth. Reanim. 24: 361-365.

Bracken, G. K. 1965. Effects of dietary components on fecundity of the parasitoid Exeristes
comstockii (Cress) (Hymenoptera: Ichneumonidae). Can. Entomol. 97: 1037-1041.









Brandl, M. T. 2006. Fitness of human enteric pathogens on plants and implications for food
safety. Annu. Rev. Phytopathol. 44: 367-392.

Breuer, T., D. H. Benkel, R. L. Shapiro, W. N. Hall, M. M. Winnett, M. J. Linn, J.
Neimann, T. J. Barrett, S. Dietrich, F. P. Downes, D. M. Toney, J. L. Pearson, H.
Rolka, L. Slutsker, P. M. Griffin, and the Investigation Team. 2001. A multistate
outbreak of Escherichia coli 0157:H7 infections linked to alfalfa sprouts grown from
contaminated seeds. Emerg. Infect. Dis. 7: 977-982.

Brichta-Harhay, D. M., M. Arthur, J. M. Bosileva, M. N. Guerini, N. Kalchayanand, M.
and Koohmaraie. 2007. Enumeration of Salmonella and Escherichia coli 0157:H7 in
ground beef, cattle carcass, hide and faecal samples using direct plating methods. J. Appl.
Microbiol. 103: 1657-1668.

Broce, A. B. 1988. An improved alsynite trap for stable flies, Stomoxys calcitrans (Diptera:
Muscidae). J. Med. Entomol. 25: 406-409.

Broce, A. B. 1993a. Dispersal of house flies and stable flies. pp. 61-69. In G. D. Thomas and S.
R. Skoda [eds.]. Rural flies in the urban environment? North Central Regional Res. Bull.
335, Institute of Agriculture and Natural Resources Res. Bull. 317. University of
Nebraska Agricultural Research Division, Lincoln, NE.

Broce, A. B. 1993b. Electrocuting and electronic insect traps: trapping efficiency and production
of airborne particles. Int. J. Environ. Health Res. 3: 47-58.

Broce, A. B. and R. J. Elzinga. 1984. Comparison of prestomal teeth in the face fly (Musca
autumnalis) and the house fly (Musca domestic) (Diptera: Muscidae). J. Med. Entomol.
21: 82-85.

Broce, A. B. and J. E. Urban. 1998. Potential microbial health hazards associated with
operation of bug zappers. Ann. Meeting Am. Soc. Microbiol. Q-252.

Buma, R., T. Maeda, Y. Marutaka, M. Kamei, H. Nagamune, and H. Kourai. 2004.
Vectorial capacity of larvae, pupae and adult of housefly (Musca domestic) for
Escherichia coli 0157:H7 and the possibility of transmission from source to human.
Med. Entomol. Zool. 55: 95-106.

Buma, R., H. Sanada, T. Maeda, M. Kamei, and H. Kourai. 1999. Isolation and
characterization of pathogenic bacteria, including Escherichia coli 0157:H7, from flies
collected at a dairy farm field. Med. Entomol. Zool. 50: 313-321.

Burg, J. G. and R. C. Axtell. 1984. Monitoring house fly, Musca domestic (Diptera:
Muscidae), populations in caged-layer poultry houses using a baited jug-trap. Environ.
Entomol. 13: 1083-1090.









Butler, J., A. Garcia-Maruniak, F. Meek, and J. E. Maruniak. 2010. Wild Florida house flies
(Musca domestic L.) as carriers of pathogenic bacteria. Fla. Entomol. 93: 218-223.

Buzby, J. C., T. Roberts, C. T. Jordan Lin, and J. M. MacDonald. 1996. Bacterial foodborne
disease: medical costs and productivity losses. Agr. Econ. Rep. No. AER741, August
1996.

Byrd, J. H. and J. L. Castner. 2009. Forensic entomology: the utility of arthropods in legal
investigations, 2nd ed., CRC Press, Boca Raton, FL.

Cafarchia, C., R. P. Lia, D. Romito, and D. Otranto. 2009. Competence of the housefly,
Musca domestic, as a vector ofMicrosporum canis under experimental conditions. Med.
Vet. Entomol. 23: 21-25.

Cagney, C., H. Crowley, G. Duffy, J. J. Sheridan, S. O'Brien, E. Carney, W. Anderson, D.
A. McDowell, I. S. Blair, and R. H. Bishop. 2004. Prevalence and numbers of
Escherichia coli 0157:H7 in minced beef and beef burgers from butcher shops and
supermarkets in the Republic of Ireland. Food Microbiol. 21: 203-212.

Callaway, T. R., R. C. Anderson, T. S. Edrington, K. J. Genovese, K. M. Bischoff, T. L.
Poole, Y. S. Jung, R. B. Harvey, and D. J. Nisbet. 2004. What are we doing about
Escherichia coli 0157:H7 in cattle? J. Anim. Sci. 82: E93-E99.

Callaway, T. R., R. O. Elder, J. E. Keen, R. C. Anderson, and D. J. Nisbet. 2003. Forage
feeding to reduce preharvest Escherichia coli populations in cattle, a review. J. Dairy Sci.
86: 852-860.

Caprioli, A., S. Morabito, H. BrugEre, and E. Oswald. 2005. Enterohaemorrhagic Escherichia
coli: emerging issues on virulence and modes of transmission. Vet. Res. 36: 289-311.

Carlson, D. A., U. R. Bernier, J. A. Hogsette, and B. D. Sutton. 2001. Distinctive
hydrocarbons of the black dump fly, Hydrotaea aenescens (Diptera: Muscidae). Arch.
Insect Biochem. Physiol. 48: 167-178.

[CDC] Centers for Disease Control and Prevention. 1983. International notes: outbreak of
hemorrhagic colitis -- Ottawa, Canada. Morb. Mort. Wkly. Rep. 32: 133-134.
(http://www.cdc.gov/mmwr/preview/mmwrhtml/00001271.htm).

[CDC] Centers for Disease Control and Prevention. 1994. E. coli 0157:H7: procedure for
isolation and identification from stool specimens. (http://wonder.cdc.gov/wonder/
PrevGuid/p0000445/P0000445. asp).

[CDC] Centers for Disease Control and Prevention. 2008. Shiga toxin-producing Escherichia
coli (STEC) case surveillance. (http://www.cdc.gov/outbreaks.html).









[CDC] Centers for Disease Control and Prevention. 2009. Multistate outbreak ofE. coli
057:H7 infections associated with beef from JBS Swift Beef Company.
(http://www.cdc.gov/ecoli/2009/0701.html).

Cebula, T. A., W. L. Payne, and P. Feng. 1995. Simultaneous identification of strains of
Escherichia coli serotype 0157:H7 and their Shiga-like toxin type by mismatch
amplification mutation assay-multiplex PCR. J. Clin. Microbiol. 33: 248-250.

Chapin, J. B. 1989. Common names of insects. Bull. Entomol. Soc. Am. 35: 177-180.

Chapman, P. A. and C. A. Siddons. 1996. A comparison of immunomagnetic separation and
direct culture for the isolation of verocytotoxin-producing Escherichia coli 0157 from
cases of bloody diarrhoea, non-bloody diarrhoea and asymptomatic contacts. J. Med.
Microbiol. 44: 267-271.

Chapman, P. A., C. A. Siddons, A. T. Cerdan Malo, and M. A. Harkin. 1997. A 1-year study
ofEscherichia coli 0157 in cattle, sheep, pigs and poultry. Epidemiol. Infect. 119: 245-
250.

Chapman, P. A., C. A. Siddons, D. J. Wright, P. Norman, J. Fox, and E. Crick. 1993. Cattle
as a possible source of verocytotoxin-producing Escherichia coli 0157 infections in man.
Epidemiol. Infect. 111: 439-447.

Chapman, P. A., D. J. Wright, and C. A. Siddons. 1994. A comparison of immunomagnetic
separation and direct culture for the isolation of verocytotoxin-producing Escherichia coli
0157 from cases bovine faeces. J. Med. Microbiol. 40: 424-427.

Chou, C.-C. and S.-J. Cheng. 2000. Recovery of low-temperature stressed E. coli 0157:H7 and
its susceptibility to crystal violet, bile salt, sodium chloride and ethanol. Int. J. Food
Microbiol. 61: 127-136.

Christensen, C. M. 1982. External parasites of dairy cattle. J. Dairy Sci. 65: 1289-2193.

Cirillo, V. J. 2006. Winged sponges: Houseflies as carriers of typhoid fever in 19th- and early
20th-century military camps. Perspect. Biol. Med. 49: 52-63.

Cohen, D., M. Green, C. Block, R. Slepon, R. Ambar, S. S. Wasserman, and M. M. Levine.
1991. Reduction of transmission of shigellosis by control of houseflies (Musca
domestica. Lancet. 337: 993-997.

Conn, D. B., J. Weaver, L. Tamang, and T. K. Craczyk. 2007. Synanthropic flies as vectors
of Cryptosporidium and Giardia among livestock and dairy-collected life in a
multispecies agricultural complex. Vector Borne Zoonotic Dis. 7: 63-652.









Cornick, N. A., S. L. Booher, T. A. Casey, and H. W. Moon. 2000. Persistent colonization of
sheep by Escherichia coli 0157:H7 and other E. coli pathotypes. Appl. Environ.
Microbiol. 66: 4926-4934.

Couteau, D., A. L. McCartney, G. R. Gibson, G. Williamson, and C. B. Faulds. 2001.
Isolation and characterization of human colonic bacteria able to hydrolyse chlorogenic
acid. J. Appl. Microbiol. 90: 873-881.

Crespo, D. C., R. E. Lecuona, and J. A. Hogsette. 1998. Biological control: an important
component in integrated management of Musca domestic (Diptera: Muscidae) in caged-
layer poultry houses in Buenos Aires, Argentina. Biol. Control 13: 16-24.

[CSTE] Council of State and Territorial Epidemiologists. 2005. Revision of the
enterohemorrhagic Escherichia coli (EHEC) condition name to Shiga toxin-producing
Escherichia coli (STEC) and adoption of serotype specific national reporting for STEC.
Position Statement 05-ID-07. (http://www.Cste.Org/Position%20statements/
Searchbyyear2005).

Davis, M. A. 2006. Comparison of cultures from rectoanal-junction mucosal swabs and feces for
detection of Escherichia coli 0157 in dairy heifers. Appl. Environ. Microbiol. 72: 3766-
3770.

Davis, M. A., K. A. Cloud-Hansen, J. Carpenter, and C. J. Hovde. 2005. Escherichia coli
0157:H7 in environments of culture-positive cattle. Appl. Environ. Microbiol. 71: 6816-
6822.

De Jesus, Antonio J.; Olsen, Alan R.; Bryce, John R., and Whiting, Richard C. 2004.
Quantitative contamination and transfer of Escherichia coli from foods by houseflies,
Musca domestic L. (Diptera: Muscidae). Int. J. Food Microbiol. 93: 259-262.

de la Torre-Bueno, J. R., S. W. Nichols, G. S. Tulloch, and R. T. Schuh. 1989. The Torre-
Bueno glossary of entomology. New York Entomological Society, New York, NY.

Delthier, I. R. 1976. The hungry fly. Harvard University Press, Cambridge, MA.

Denholm, I., R. M. Sawicki, and A. W. Farnham. 1985. Factors affecting resistance to
insecticides in house flies, Musca domestic L. (Diptera:Muscidae). IV. The population
biology of flies on animal farms in south eastern England and its implications for the
management of resistance. Bull. Entomol. Res. 1985; 75: 144-158.

Denny, J., M. Bhat, and K. Eckmann. 2008. Outbreak of Escherichia coli 0157:H7 associated
with raw milk Consumption in the Pacific Northwest. Foodborne Path. Dis. 5: 321-328.

Desmarchelier, P. M., S. S. Bilge, N. Fegan, L. Mills, J. C. Vary Jr., and P. I. Tarr. 1998. A
PCR specific for Escherichia coli 0157 based on the rfb locus encoding 0157
lipopolysaccharide. J. Clin. Microbiol. 36: 1801-1804.









Dhillon, M. S., and G. L. Challet. 1985. The evaluation of three sampling techniques for the
determination of fly (Diptera) densities at four sanitary landfills in southern California.
Bull. Soc. Vect. Ecol. 10: 36-40.

Doane, C. A., P. Pangloli, H. A. Richards, J. R. Mount, D. A. Golden, and F. A. Draughon.
2002. Occurrence ofEscherichia coli 0157:H7 in diverse farm environments. J. Food
Prot. 70: 6-10.

Donnenberg, M. S. 2002. Introduction. pp. xxi-xxv. In Donnenberg, M. S. [ed.]. Escherichia
coli: Virulence mechanisms of a versatile pathogen. Academic Press, San Diego, CA.

Donnenberg, M. S. and T. S. Whittam. 2002. Pathogenesis and evolution of virulence in
enteropathogenic and enterohemorrhagic Escherichia coli. J. Clin. Investigation. 107:
539-548.

Dougherty, E. C. 1959. Introduction of axenic culture of invertebrate metazoa: a goal. Ann. N.
Y. Acad. Sci. 77: 27-54.

Doyle, M. E., J. Archer, C. W. Kaspar, and R. Weiss. 2006. FRI Briefings: Human illness
caused by E. coli 0157:H7 from food and non-food sources. Food Research Institute,
University of Wisconsin-Madison, Madison, WI.

Doyle, M. P. and L. R. Beuchat. 2007. Food microbiology: fundamentals and frontiers. ASM
Press, Washington, D.C.

Doyle, M. P. and J. L. Schoeni. 1984. Survival and growth characteristics of Escherichia coli
associated with hemorrhagic colitis. Appl. Environ. Microbiol. 48: 855-856.

Duffy, G. 2003. Verocytoxigenic Escherichia coli in animal faeces, manures and slurries. J.
Appl. Microbiol. 94: 94S-103S.

Dunn, J. R., J. E. Keen, D. Moreland, and R. A. Thompson. 2004a. Prevalence of
Escherichia coli 0157:H7 in white-tailed deer from Louisiana. J. Wildlife Dis. 40: 361-
365.

Dunn, J. R., J. E. Keen, and R. A. Thompson. 2004b. Prevalence of shiga-toxigenic
Escherichia coli 0157:H7 in adult dairy cattle. J. Am. Vet. Med. Assoc. 224: 1151-1158.
DuPont, H. L. 2007. The growing threat of foodborne bacterial enteropathogens of animal
origin. Clin. Infect. Dis. 45: 1353-1361.

Duriez, P., Y. Zhang, Z. Lu, A. Scott, and E. Topp. 2008. Loss of virulence genes in
Escherichia coli populations during manure storage on a commercial swine farm. Appl.
Environ. Microbiol. 74: 3935-3942.









Durso, L. M. and J. E. Keen. 2007. Shiga-toxigenic Escherichia coli 0157 and non-Shiga-
toxigenic E. coli 0157 respond differently to culture and isolation from naturally
contaminated bovine faeces. J. Appl. Microbiol. 103: 2457-2464.

Dynal. 2007. Dynabeads anti-E. coli 0157 Manual. Invitrogen Dynal AS. Oslo, Norway.

Echeverria, P., B. A. Harrison, C. Tirapat, and A. McFarland. 1983. Flies as a source of
enteric pathogens in a rural village in Thailand. Appl. Environ. Microbiol. 46: 32-36.

Echeverry, A., G. H. Loneragan, B. A. Wagner, and M. M. Brashears. 2005. Effect of
intensity of fecal pat sampling on estimates of Escherichia coli 0157 prevalence. Am. J.
Vet. Res. 66: 2023-2027.

Eddy, G. W., A. R. Roth, and F. W. Plapp, Jr. 1962. Studies on the flight habits of some
marked insects. J. Econ. Entomol. 55: 603-608.

[ESA] Entomological Society of America. 2009. ESA common names of insects and related
organisms online database. (http://www.entsoc.org/Pubs/CommonNames/ search.asp.
Accessed 11 January 2008).

Euzeby, J. P. M. 2008. List of prokaryotic names with standing in nomenclature (LPSN):
formerly, list of bacterial names with standing in nomenclature (LBSN).
(http://www.bacterio.cict.fr/aldl.html).

[Excel] Microsoft Excel. 2003. Microsoft Excel User's Manual, Ver. 11. Microsoft Corp.,
Redmond, WA.

[FDA-CFSAN] Food and Drug Administration, Center for Food Safety and Applied
Nutrition. 2007a. Media index for BAM. Bacteriological Analytical Manual (BAM)
Online. (http://www.fda.gov/Food/ScienceResearch/LaboratoryMethods/
BacteriologicalAnalyticalManualBAM/ucm055778.htm).

[FDA-CFSAN] Food and Drug Administration, Center for Food Safety and Applied
Nutrition. 2007b. Diarrheagenic Escherichia coli. Bacteriological Analytical Manual
(BAM) Online. (http://www.cfsan.fda.gov/-ebam/bam-4a.html 10 p).

[FDA-CFSAN] Food and Drug Administration, Center for Food Safety and Applied
Nutrition. 2007c. Escherichia coli 0157:H7. Foodborne pathogenic microorganisms and
natural toxins handbook: the "bad bug book." (http://www.cfsan.fda.gov/-mow/
chap 15.html).

Feder, I., F. M. Wallace, J. T. Gray P. Fratamico, P. J. Fedorka-Cray, R. A. Pearce, J. E.
Call, R. Perrine, and J. B. Luchansky. 2003. Isolation of Escherichia coli 0157:H7
from intact colon fecal samples of swine. Emerg. Infect. Dis. 9: 380-383.









Feng, P. C. H. and S. R. Monday. 2005. Multiplex PCR for specific identification of
enterohemorrhagic Escherichia coli strains in the 0157:H7 complex. In C. C. Adley
[ed.]. Methods in Biotech., Vol. 21: Food-borne pathogens: methods and protocols.
Humana Press Inc, Totowa, NJ.

Floate, K. D., P. Coghlin, and G. A. P. Gibson. 2000. Dispersal of the filth fly parasitoid
Muscidifurax raptorellus (Hymenoptera: Pteromalidae) following mass releases in cattle
confinements. Biol. Control: 172-178.

Floyd, T. M. and B. H. Cook. 1954. The housefly as a carrier of pathogenic human enteric
bacteria in Cairo. J. Egypt. Publ. Health Assoc. 28: 75-85.

Fode-Vaughn, K. A., J S. Maki, J. A. Benson, and M. L. P. Collins. 2003. Direct PCR
detection of Escherichia coli 0157:H7. Lett. Appl. Microbiol. 37: 239-243.

[Food Consumer] Food Consumer. 2006. Nationwide E. coli 0157:H7 outbreak: questions and
answers. (http://www.foodconsumer.org/777/8/Nationwide _EColi_O157 H7
OutbreakQuestions_ampAnswers.shtml).

Foster, S. P. and M. O. Harris. 1997. Behavioral manipulation methods for insect pest-
management. Annu. Rev. Entomol. 42: 123-146.

Fotedar, R., U. Banerjee, S. Singh, Shriniwas, and A. K. Verma. 1992. The housefly (Musca
domestic) as a carrier of pathogenic microorganisms in a hospital environment. J. Hosp.
Infect. 20: 209-215.

Frandson, R. D. 1969. Anatomy and physiology of farm animals. Lea and Febiger, Philadelphia,
PA.
Fratamico, P. M. and D. O. Bayles. 2005. Molecular approaches for detection, identification,
and analysis of foodborne pathogens, pp. 1-14. In A. K. Bhunia, and J. L. Smith (eds.).
Foodborne pathogens. Caister Acad. Press, Norfolk, UK.

Fratamico, P. M., S. K. Sackitey, M. Wiedmann, and M. Y. Deng. 1995. Detection of
Escherichia coli 0157:H7 by multiplex PCR. J. Clin. Microbiol. 33: 2188-2191.

Fremaux, B., C. Prigent-Combaret, and C. Vernozy-Rozand. 2008. Long-term survival of
Shiga toxin-producing Escherichia coli in cattle effluents and environment: An updated
review. Vet. Microbiol. 132: 1-18.

Frenzen, P. D., A. Drake, and F. J. Angulo. 2005. Economic cost of illness due to Escherichia
coli 0157 infections in the United States. J. Food Prot. 68: 2623-2630.

Frick, T. B. and D. W. Tallamy. 1996. Density and diversity of nontarget insects killed by
suburban electric insect traps. Entomol. News 107: 77-82.









Fried, J. H., D. J. Levey, and J. A. Hogsette. 2005. Habitat corridors function as both drift
fences and movement conduits for dispersing flies. Oecologia 143: 645-651.

Galland, J. C., D. R. Hyatt, S. S. Crupper, and D. W. Acheson. 2001. Prevalence, antibiotic
susceptibility, and diversity of Escherichia coli 0157:H7 isolates from a longitudinal
study of beef cattle feedlots. Appl. Environ. Microbiol. 67: 1619-1627.

Gannon, V. P. J., S. D'Souza, T. Graham, R. K. King, K. Rahn, and S. Read. 1997. Use of
the flagellar H7 gene as a target in multiplex PCR assays and improved specificity in
identification of enterohemorrhagic Escherichia coli strains. J. Clin. Microbiol. 35: 656-
662.

Geden, C. J. 2006. Visual targets for capture and management of house flies, Musca domestic
L. J. Vector Ecol. 31: 152-157.

Geden, C. J., J. A. Hogsette, and R. D. Jacobs. 1999. Effect of airflow on house fly (Diptera:
Muscidae) distribution in poultry houses. J. Econ. Entomol. 92: 416-20.

Geden, C. J., D. C. Steinkraus, D. A. Rutz. 1993. Evaluation of two methods for release of
Entomophthora muscae (Entomophthorales: Entomophthoraceae) to infect house flies
(Diptera: Muscidae) on dairy farms. Environ. Entomol. 20: 1201-1208.

[Genvault] GenVault Corp. 2010. DNA Quantitation: methods and recommendations in use at
GenVault. (www.genvault.com).

Gillespie, J. R. 2002. Modern livestock and poultry production, 6th ed. Thomas Delmare
Learning, Albany, NY.

Google Earth. 2009. Google Earth. (http://earth.google.com/download-earth.html).

Goulson, D., L. C. Derwent, M. E. Hanley, D. W. Dunn, and S. R. Abolins. 2005. Predicting
calyptrate fly populations from the weather, and probably consequences of climate
change. J. Appl. Ecol. 42: 795-804.

Graham-Smith, G. S. 1912. An investigation into the possibility of pathogenic microorganisms
being taken up by the larva and subsequently distributed by the fly, pp. 330-335. In 41st
Ann. Rep. Local Govt. Bd. Suppl. Rep. Med. Off. 1911-1912. App. B.

Graham-Smith. 1939. .Further observations on the relation of the decline in the number of
horse-drawn vehicles to the fall in the summer diarrhoea death-rate. J. Hyg.39:558-562.

Gratz, N. G. 1999. Emerging and resurging vector-borne diseases. Annu. Rev. Entomol. 44: 51-
75.

Greenberg, B. 1959a. House fly nutrition. II. Comparative survival values of sucrose and water.
Ann. Entomol. Soc. Am. 53: 125-128.









Greenberg, B. 1959b. Persistence of bacteria in the developmental stages of the housefly. I.
Survival of enteric pathogens in the normal and aseptically reared host. Am. J. Trop.
Med. Hyg. 8:405-411.

Greenberg, B. 1959c. Persistence of bacteria in the developmental stages of the housefly. II.
Quantitative study of the host-contaminant relationship in flies breeding under natural
conditions. Am. J. Trop. Med. Hyg. 8: 412-416.

Greenberg, B. 1959d. Persistence of bacteria in the developmental stages of the housefly. III.
Quantitative distribution in prepupae and pupae. Am. J. Trop. Med. Hyg. 8: 613-617.

Greenberg, B. 1959e. Persistence of bacteria in the developmental stages of the housefly. IV.
Infectivity of the newly emerged adult. Am. J. Trop. Med. Hyg. 8: 618-22.

Greenberg, B. 1965. Flies and disease. Sci. Am. 213: 92-99.

Greenberg, B. 1971. Flies and disease, Vol. I ecology, classification and biotic associations.
Princeton University Press, Princeton, NJ.

Greenberg, B. 1973. Flies and disease, Vol. II biology and disease transmission. Princeton
University Press, Princeton, NJ.

Greenberg, B., J. A. Kowalski, and M. J. Klowden. 1970. Factors affecting the transmission of
Salmonella by flies: natural resistance to colonization and bacterial interference. Infect.
Immun. 2: 800-809.

Greenquist, M. A., J. S. Drouillard, J. M. Sargeant, B. E. Depenbusch, X. Shi, K. F.
Lechtenberg, and T. G. Nagaraja. 2005. Comparison of rectoanal mucosal swab
cultures and fecal cultures for determining prevalence of Escherichia coli 0157:H7 in
feedlot cattle. Appl. Environ. Microbiol. 71: 6431-6433.

Grif K., M. P. Dierich, H. Karch, F. Allerberger. 1998. Strainspecific differences in the
amount of Shiga toxin released from enterohaemorrhagic Escherichia coli 0157
following exposure to subinhibitory concentrations of antimicrobial agents. Eur. J. Clin.
Microbiol. Infect. Dis. 17: 761-766.

Griibel, P., J. S. Hoffman, F. K. Chong, N. A. Burstein, C. Mepani, and D. R. Cave. 1997.
Vector potential of houseflies (Musca domestic) for Helicobacterpylori. J. Clin.
Microbiol. 35: 1300-1303.

Gullan, P. J. and P. S. Cranston. 2000. The insects: an outline of entomology, 2nd ed.
Blackwell Science, Osney Mead, Oxford.

Hagler, J. R. and C. G. Jackson. 2001. Methods for marking insects: current techniques and
future prospects. Annu. Rev. Entomol. 46: 511-543.









Hald, B., H. Skovgard, D. D. Bang, K. Pedersen, J. Dybdahl, J. B. Jespersen, and M.
Madsen. 2004. Flies and Campylobacter infection of broiler flocks. Emerg. Infect. Dis.
10: 1490-1492.

Halverson, M. 2000. The price we pay for corporate hogs. Institute for Agriculture and Trade
Policy (IATP) Report. July 2000, 248 pp. (http://www.iatp.org/hogreport).

Hammer, 0. 1941. Biological and ecological investigations on flies associated with pasturing
cattle and their excrement. Vidensk. Meddr. Dansk. Naturh. Foren. 105: 5-257.

Hancock, D. D., T. E. Besser, M. L. Kinsel, P. I. Tarr, D. H. Rice, and M. G. Paros. 1994.
The prevalence ofEscherichia coli 0157:H7 in dairy and beef cattle in Washington State.
Epidemiol. Infect. 113: 199-207.

Hancock, D., D. Rice, L. Thomas, D. Dargataz, and T. Besser. 1997. Epidemiology of
Escherichia coli 0157 in feedlot cattle. J. Food Prot. 60: 462-465.

Hanec, W. 1956. A study of the environmental factors affecting dispersion of house flies (Musca
domestic L.) in a dairy community near Fort Whyte, Manitoba. Can. Entomol. 88: 270-
272.

Hatch Jr., E. 1911. The housefly as a carrier of disease. Ann. Am. Acad. Political Soc. Sci. 37:
168-179.

Haupt, A. and J. R. Busvine. 1968. The effect of overcrowding on the size of houseflies
(Musca domestic L.). Trans. R. Entomol. Soc. Lond. 120: 297-311.

Hawley, J. E., L. R. Penner, S. E. Wedberg, and W. Kulp. 1951. The role of the house fly, ii,
in the multiplication of certain enteric bacteria. Am. J. Trop. Med. Hyg. 31: 572-582.

Heuvelink, A. E. 2003. Review of media for the isolation of diarrhoeagenic Escherichia coli, pp.
229-247. In J. E. L. Corry, G. D. W. R. Curtis, and M. E. Baird [eds.]. Handbook of
culture media for food microbiology. Elsevier Science, Amsterdam.

Hewitt, C. G. 1914. The house-fly, its structure, habits, development, relation to disease control.
University Press, Cambridge, UK.

Hibbing, M. E., C. Fuqua, M. R. Parsek, and S. Brook Peterson. Bacterial competition:
surviving and thriving in the microbial jungle. Nature Rev. Microbiol. 8: 15-25.

Hoffmann, J. A. and C. Hetru. 1992. Insect defensins: inducible antibacterial peptides.
Immunol. Today 13: 411-415.

Hoffman, R. A. and A. W. Lindquist. 1951. Studies on treatment of flies with radioactive
phosphorus. J. Econ. Entomol. 44: 471-473.









Hogsette, J. A. 1983. An attractant self-marking devise for marking field populations of stable
flies with fluorescent dusts. J. Econ. Entomol. 76: 510-514.

Hogsette, J. A. 1984. Effect of flourescent dust color on the attractiveness of attractant self-
marking devises to the stable fly (Diptera: Muscidae). J. Econ. Entomol. 77: 130-132.

Hogsette, J. A. 1992. New diets for production of house flies and stable flies (Diptera:
Muscidae) in the laboratory. J. Econ. Entomol. 85: 2291-2294.

Hogsette, J. A. 1996. Development of house flies (Diptera:Muscidae) in sand containing varying
amounts of manure solids and moisture. J. Econ. Entomol. 89: 940-945.

Hogsette, J. A. 2008. Ultraviolet light traps: design affects attraction and capture. In Proc. 6th
Int. Conf. Urb. Pests, Hungary, 4 pp. OOK-Press Kft., Hungary.

Hogsette, J. A., D. A. Carlson, and A. S. Nejame. 2002. Development of granular boric acid
sugar baits for house flies (Diptera: Muscidae). J. Econ. Entomol. 95: 1110-1112.

Hogsette, J. A., R. Farkas, and C. Thur6czy. 2001. Hymenopteran pupal parasitoids recovered
from house fly and stable fly (Diptera: Muscidae) pupae collected on livestock facilities
in Southern and Eastern Hungary. Environ. Entomol. 30: 107-111.

Hogsette, J. A., and R. D. Jacobs. 1999. Failure ofHydrotaea aenescens, a larval predator of
the house fly, Musca domestic L., to establish in wet poultry manure on a commercial
farm in Florida, USA. Med. Vet. Entomol. 13: 349-354.

Hogsette, J. A., R. D. Jacobs, and R. W. Miller. 1993. The sticky card: device for studying the
distribution of adult house fly (Diptera: Muscidae) populations in closed poultry houses.
J. Econ. Entomol. 86: 450-454.

Hogsette, J. A. and J. P. Ruff. 1990. Comparative attraction of four different fiberglass traps to
various age and sex classes of stable fly (Diptera: Muscidae) adults. J. Econ. Entomol.
83: 883-886.

Hogsette, J. A. and F. Washington. 1995. Quantitative mass production of Hydrotaea
aenescens (Diptera: Muscidae). J. Econ. Entomol. 88: 1238-1242.

Holland, J. L., L. Louie, A. E. Simor, and M. Louie. 2000. PCR detection of Escherichia coli
0157:H7 directly from stools: Evaluation of commercial extraction methods for purifying
fecal DNA. J. Clin. Microbiol. 38: 4108-4113.

Holzapfel, E. P. and J. C. Harrell. 1968. Transoceanic dispersal studies of insects. Pacific
Insects 10: 115-153.









Howard, L. O. 1900. A contribution to the study of the insect fauna of human excrement (with
especial reference to the spread of typhoid fever by flies). Proc. Wash. Acad. Sci. 2: 541-
604.

Howard, L. O. 1910. The house fly, disease carrier: an account of its dangerous activities and of
the means of destroying it. Frederick A. Stokes Co. Publ. New York, NY.

Hsu, C., T. Tsai, and T. Pan. 2005. Use of the duplex TaqMan PCR system for detection of
Shiga-like toxin-producing Escherichia coli 0157. J. Clin. Microbiol. 43: 2668-2673.

Hu, Y., Q. Zhang, and J. C. Meitzler. 1999. Rapid and sensitive detection of Escherichia coli
0157:H7 in bovine faeces by a multiplex PCR. J. Appl. Microbiol. 87: 867-876.

Hussein, H. S. and T. Sakuma. 2005. Prevalence of Shiga toxin-producing Escherichia coli in
dairy cattle and their products. J. Dairy Sci. 88: 450-466.

Hussein, H. S., B. H. Thran, M. R. Hall, and W. G. Kvasnicka. 2003. Verotoxin-producing
Escherichia coli in culled beef cows grazing rangeland forages. Exp. Biol. Med. 228:
352-357.

Hutchison, R. H. 1916. Notes on the preoviposition of the house fly, Musca domestic L. U. S.
Dept. Agr. Bull., Washington, D. C. 345: 1-16.

[IOWH] Institute for One World Health. 2008. Diarrheal diseases fact sheet,
(http://www.oneworldhealth.org/diseases/diarrhea.php).

Islam, M., M. P. Doyle, S. C. Phatak, P. Millner, and X. Jiang. 2004. Persistence of
enterohemorrhagic Escherichia coli 0157:H7 in soil and on leaf lettuce and parsley
grown in fields treated with contaminated manure composts or irrigation water. J. Food
Prot. 67: 1365-1370.

Islam, M., M. P. Doyle, S. C. Phatak P. Millner, and X. Jiang. 2005. Survival of Escherichia
coli 0157:H7 in soil and on carrots and onions grown in fields treated with contaminated
manure composts or irrigation water. Food Microbiol. 22: 63-70.

[ITIS] Integrated Taxonomic Information System. 2008. Musca domestic Linnaeus, 1758.
ITIS Report. (http://www.itis.gov).

Janda, J. M. and S. L. Abbott. 2006. The Enterobacteria, 2nd edition. ASM Press, Washington,
D.C.

Jiang, X., J. Morgan and M. P. Doyle. 2002.Fate of Escherichia coli 0157:H7 in manure-
amended soil. Appl. Environ. Microbiol. 68: 2605-2609.

Johnson, C. G. 1966. A functional system of adaptive dispersal of flight. Annu. Rev. Entomol.
11: 233-260.









Johnson, C. G. 1969. Migration and dispersal of insects by flight. Methuen and Co., London,
UK.

Johnson, J. R. 2002. Evolution of pathogenic Escherichia coli. pp. 55-77. In M. S. Donnenberg
[ed.], Escherichia coli: Virulence mechanisms of a versatile pathogen. Academic Press,
San Diego, CA.

Johnson, R. P., J. B. Wilson, P. Michel, K. Rahn, S. A. Renwick, C. L. Gyles, and J. S.
Spika. 1999. Escherichia coli 0157 in farm animals. CABI Publishing, Wallingford,
Oxon, UK.

Jones, C. J., J. A. Hogsette, R. S. Patterson, D. E. Milne, G. D. Propp, J. F. Milio,, L G.
Rickard, and J. P. Ruff. 1991. Origin of stable flies (Diptera: Muscidae) on West
Florida beaches: electrophoretic analysis of dispersal. J. Med. Entomol. 28: 787-795.

Karch, H., C. Janetzke-Mittmann, S. Aleksic, and M. Datz. 1996. Isolation of
enterohemorrhagic Escherichia coli 0157 strains from patients with hemolytic-uremic
syndrome by using immunomagnetic separation, DNA-based methods, and direct culture.
J. Clin. Microbiol. 34: 516-519.

Karmali, M. 1989. Infection by verocytotoxin-producing Escherichia coli. Clin. Microbiol. Rev.
2: 15-38.

Kaufman, P. E. 2002. Dairy pest management, arthropods, pp. 181-183. In D. Pimentel [Ed.],
Encyclopedia of pest management, Vol. 1. Marcel Dekker, Inc.

Kaufman, P. E., M. Burgess and D. A. Rutz. 2002. Population dynamics of manure inhabiting
arthropods under an integrated pest management (IPM) program in New York poultry
program in New York poultry facilities 3 case studies. J. Appl. Poul. Sci. Res. 11: 90-
103.

Kaufman, P. E., S. J. Long, and D. A. Rutz. 2001a. Impact of exposure length and pupal
source on Muscidifurax raptorellus and Nasonia vitripennis (Hymenoptera:
Pteromalidae) parasitism in a New York poultry facility. J. Econ. Entomol. 94: 998-1003.

Kaufman, P. E., S. J. Long, D. A. Rutz, and J. K. Waldron. 2001b. Parasitism rates of
Muscidifurax raptorellus and Nasonia vitripennis (Hymenoptera: Ptermolidae) after
individual and paried releases in New York poultry facilities. J. Econ. Entomol. 94: 593-
598.

Kaufman, P. E., C. Reasor, D. A. Rutz, J. K. Ketzis, and J. J. Ahrends. 2005a. Evaluation of
Beauveria bassiana applications against adult house flies, Musca domestic, in
commercial caged-layer poultry facilities in New York state. Biol. Control 33: 360-367.

Kaufman, P. E., D. A. Rutz, and S. Frisch. 2005b. Large sticky traps for capturing house flies
and stable flies in dairy calf greenhouse facilities. J. Dairy Sci. 88: 176-181.









Kaufman, P. E., J. G. Scott, and D. A. Rutz. 2001. Monitoring insecticide resistance in house
flies (Diptera: Muscidae) from New York dairies. Pest Manag. Sci. 57: 514-521.

Keen, J. E., T. F.Wittum, J. R. Dunn, J. L. Bono, and L. M. Durso. 2006. Shiga-toxigenic
Escherichia coli 0157 in agricultural fair livestock, United States. Emerg. Infect. Dis. 12:
780-786.

Klein, E. J., J. R. Stapp, M. A. Neill, J. M. Besser, M. T. Osterholm, and P. I. Tarr. 2004.
Shiga toxin antigen detection should not replace Sorbitol MacConkey agar screening of
stool specimens. J. Clin. Microbiol. 42: 4416-4417.

Kobayashi, M., T. Sasaki, N. Saito, K. Tamur, K. Suzuki, H. Watanabe, and N. Agui. 1999.
Houseflies: not simple mechanical vectors of enterohemorrhagic Escherichia coli
0157:H7. Am. J. Trop. Med. Hyg. 61: 625-629.

Kosek, M., C. Bern, and R. L. Guerrant. 2003. The global burden of diarrhoeal disease, as
estimated from studies published between 1992 and 2000. Bull. World Health Organ. 81:
197-204.

Kovacs Sr., F., I. Medveczky, L. Papp and E. Gondar. 1990. Role of prestomal teeth in
feeding of the house fly, Musca domestic (Diptera; Muscidae). Med. Vet. Entomol. 4:
331-335.

Krafsur, E. S., M. A. Cummings, M. A. Endsley, J. G. Marquez, and J. D. Nason. 2005.
Geographic differentiation in the house fly estimated by microsatellite and mitochondria
variation. J. Hered. 96: 502-512.

Kristiansen, K. and 0. Slovmand. 1985. A method for the study of population size and survival
rate of houseflies. Entomol. Exp. Appl. 38: 145-150.

Kriiger, A., P. M. A. Lucchesi, and A. E. Parma. 2007. Evaluation of vt2-subtyping methods
for identifying vt2g in verotoxigenic Escherichia coli. J. Med. Microbiol. 56: 1474-1478.

Kudva, I. T., K. Blanch, and C. J. Hovde. 1998. Analysis of Escherichia coli 0157:H7
survival in ovine or bovine manure and manure slurry. Appl. Environ. Microbiol. 64:
3166-3174.

Kumar, M. and G. G. Carmichael. 1998. Antisense RNA: Function and fate of duplex RNA in
cells of higher eukaryotes. Microbiol. Mol. Biol. Rev. 62: 1415-1434.

Lahti, E., O. Ruoho, L. Rantala, M. Hanninen, and T.Honkanen-Buzalski. 2003.
Longitudinal study ofEscherichia coli 0157 in a cattle finishing unit. Appl. Environ.
Microbiol. 69: 554-561.









Lazarus, W., D. A. Rutz, R. W. Miller, and D. A. Brown. 1989. Costs of existing and
recommended manure management practices for house fly and stable fly (Diptera:
Muscidae) control on dairy farms. J. Econ. Entomol. 82: 1145-1151.

Ledingham, J. C. G. 1911. On the survival of specific microorganisms in pupae and imagines of
Musca domestic raised from experimentally infected larvae. Experiments with B.
typhosus. J. Hyg. 11: 333-340.

LeJeune, J. T., D. D. Hancock, and T. E. Besser. 2006. Sensitivity of Escherichia coli 0157
detection in bovine feces assessed by broth enrichment followed by immunomagnetic
separation and direct plating methodologies. J. Clin. Microbiol. 44: 872-875.

Levin, S. A. and V. Andreasen. 1999. Disease transmission dynamics and the evolution of
antibiotic resistance in hospitals and communal settings. Proc. Natl. Acad. Sci. 96: 800-
801.

Lole, M. J. 2005. Nuisance flies and landfill activities: an investigation at a West Midlands
landfill site. Waste Manag. Res. 23: 420-428.

Lysyk, T. J. 1993. Seasonal abundance of stable flies and house flies (Diptera: Muscidae) in
dairies in Alberta, Canada. J. Med. Entomol. 30: 888-895.

Lysyk, T. J. and R. C. Axtell. 1985. Comparison of baited jug-trap and spot cards for sampling
house fly, Musca domestic (Diptera: Muscidae), populations in poultry houses. Environ.
Entomol. 14: 815-819.

Lysyk, T. J. and R. C. Axtell. 1986. Field evaluation of three methods for monitoring
populations of house flies (Musca domestic) (Diptera: Muscidae) and other filth flies in
three types of poultry housing systems. J. Econ. Entomol. 79: 144-151.

Lysyk, T. J., L. D. Kalischuk-Tymensen, L. B. Selinger, R. C., Lancaster, L. Wever, and
K.-J. Cheng. 1999. Rearing stable fly larvae (Diptera: Muscidae) on an egg yolk
medium. J. Med. Entomol. 36: 382-388.

MacLeod, J. and J. Donnelly. 1960. Natural features and blowfly movement. J. Anim. Ecol. 29:
85-93.
Macloskie, G. 1880. The proboscis of the house-fly. Am. Natur. 14: 153-161.

Macovei, L. and L. Zurek. 2007. Influx of enterococci and associated antibiotic resistance and
virulence genes from ready-to-eat food to the human digestive tract. Appl. Environ.
Microbiol. 73: 6740-6747.

Macovei, L., B. Miles, and L. Zurek. 2008. Potential of houseflies to contaminate ready-to-eat
food with antibiotic-resistant enterococci. J. Food Prot. 71: 435-439.









Madigan, M. T. and J. M. Martinko. 2006. Brock biology of microorganisms. Benjamin
Cummings, San Francisco, CA.

Mai, V., C. R. Braden, J. Heckendorf, B. Pironis, and J. M. Hirshon. 2006. Monitoring of
stool microbiota in subjects with diarrhea indicates distortions in composition. J. Clin.
Microbiol. 44: 4550-4552.

Majalija, S., H. Segal, F. Ejobi, and B. G. Elisha. 2008. Shiga toxin gene-containing
Escherichia coli from cattle and diarrheic children in the pastoral systems of
southwestern Uganda. J. Clin. Microbiol. 46: 352-354.

Malik, A., N. Singh, and S. Satya. 2007. House fly (Musca domestica: A review of control
strategies for a challenging pest. J. Environ. Sci. Health, Part B 42: 453-469.

Manafi, M. 2003. Media for detection and enumeration of 'total' Enterobacteriaceae, coliforms
and Escherichia coli from water and foods, Handbook of culture media for food
microbiology. Elsevier Science, Amsterdam.

Matthews, L., J. C. Low, D. L. Gaily, M. C. Pearce, D. J. Mellor, J. A. P. Heesterbeek, M.
Chase-Topping, S. W. Naylor, D. J. Shaw, S. W. J. Reid, G. J. Gunn, and M. E. J.
Woolhouse. 2005. Heterogeneous shedding of Escherichia coli 0157 in cattle and its
implications for control. Proc. Natl. Acad. Sci. 103: 547-552.

Matthysse, J. G. 1945. Observations on housefly overwintering. J. Econ. Entomol. 38: 493-494.

McKay, T., C. D. Steelman, S. M. Brazil, and A. L. Szalanski. 2007. Sustained mass release
of pupal parasitoids (Hymenoptera: Pteromalidae) for control of Hydrotaea aenescens
and Musca domestic (Diptera: Muscidae) in broiler-breeder poultry houses in Arkansas.
J. Agr. Urban Entomol. 24: 67-85.

Mead, P. S. and P. M. Griffin. 1998. Escherichia coli 0157:H7. Lancet 352: 1207-1212.

Mead, P. S., L. Slutsker, V. Dietz, L. F. McCraig, J. S. Bresee, C. Shapiro, P. M. Griffin,
and R. V. Tauxe. 1999. Food-related illness and death in the United States. Emerg.
Infect. Dis. 5: 607-625.

[MEDIC] Medical Education Information Center. 1995. Department of Pathology and
Laboratory Medicine's. Enterobacteriaceae. (http://medic.med.uth.tmc.edu/
path/00001500.htm).

Meerburg, B. G., H. M. Vermeer, and A. Kijlstra. 2007. Controlling risks of pathogen
transmission by flies on organic pig farms. Outlook Agric. 36: 193-197.

Mennigmann, H. D. 1979. Storage death at low temperature (-18 C) of strains of Escherichia
coli with different repair capacities. J. Gen. Microbiol. 112: 207-210.









Miles, E. J. 1959. Disinfestation and control of pests on refuse tips. J. Roy. Soc. Prom. Health
79: 268-273.

Milio, J., C. S. Lofgren, and D. F. Williams. 1988. Nuptial flight studies of field-collected
colonies of Solenopsis invicta Buren, pp. 419-431. In J. C. Trager [ed.], Advances in
Myrmecology, E. J. Brill, Leiden, NY.

Moon, R. D. 2002. Muscid flies (Muscidae), pp. 279-302. In Mullen, G. and L. Durden
[eds.].Medical and veterinary entomology. Elsevier Science Academic Press, London,
UK.

Moon, R. D. and H. J. Meyer. 1985. Nonbiting flies, pp. 65-82. In Williams, R. E., R. D. Hall,
A. B. Broce, and P. J. Scholl [eds.]. Livestock entomology. John Wiley and Sons, New
York, NY.

Morabito, S., G. Dell'Omo, U. Agrimi, H. Schmidt, H. Karch, T. Cheasty, and A. Caprioli.
2001. Detection and characterization of Shiga toxin-producing Escherichia coli in feral
pigeons. Vet. Microbiol. 82: 275-283.

Morgan, N. 0. and L. G. Pickens. 1978. XI. House flies and other nonbiting flies (Family
Muscidae). United States Department of Agriculture Handbook: Surveillance and
collection of arthropods of veterinary importance 518: 72-76.

Morgan, P. B., D. E. Weidhaas, and G. C. LaBrecque. 1979. Host-parasite relationship of the
house fly, Musca domestic L., and the microhymenopteran pupal parasite, Muscidifurax
raptor Girault and Sanders (Diptera: Muscidae and Hymenoptera: Pteromalidae). J.
Kansas Entomol. Soc. 52: 276-281.

Mullen, G. and L. Durden. 2002. Medical and veterinary entomology. Elsevier Science
Academic Press, London, UK.

Murinda, S. E. and S. P. Oliver. 2006. Physiologic and molecular markers for detection of
Shiga toxin-producing Escherichia coli serotype 026 strains. Foodborne Path. Dis. 3:
163-177.

Murvosh, C. M., R. L. Fye, and G. C. Labrecque. 1964. Studies on the mating behavior of the
house fly, Musca domestic L. Ohio J. Sci. 4: 264-271.

Murvosh, C. M. and C. W. Thaggard. 1966. Ecological studies of the house fly. Ann.
Entomol. Soc. Am. 59: 533-547.

Nash, J. T. C. 1909. House flies as carriers of disease. J. Hyg. 9: 141-169.

Nash, J. T. C. 1913. Range of flight of Musca domestic. Lancet 182:1585-1586.









Nataro, J. P. and J. B. Kaper. 1998. Diarrheagenic Escherichia coli. Clin. Microbiol. Rev. 11:
132-201.

Nation, J. L. 2002. Insect physiology and biochemistry. CRC Press, Boca Raton, FL.

Nayduch, D., A. Honko, G. P. Noblet, and F. Stutzenberger. 2001. Detection ofAeromonas
caviae in the common housefly Musca domestic by culture and polymerase chain
reaction. Epidemiol. Infect. 127: 561-566.

Nayduch, D., G. P. Noblet, and F. J. Stutzenberger. 2005. Fate of bacteria, Aeromonas caviae,
in the midgut of the housefly, Musca domestic. Invert. Biol. 124: 74-78.

Naylor, S. W., J. C. Low, T. E. Besser, A. Mahajan, G. J. Gunn, M. C. Pearce, I. J.
McKendrick, D. G. E. Smith, and D. L. Gaily. 2003. Lymphoid follicle-dense mucosa
at the terminal rectum is the principal site of colonization of enterohemorrhagic
Escherichia coli 0157:H7 in the bovine host. Infect. Immun. 71: 1505-1512.

Nazni, W. A., H. Luke, W. M. Wan Rozita, A. G. Abdullah, I. Sa'diyah, A. H. Azahari, I.
Zamree, S. B. Tan, H. L. Lee, and M. A. Sofian. 2005. Determination of the flight
range and dispersal of the house fly, Musca domestic (L.) using mark release recapture
technique. Trop. Biomed. 22: 53-61.

[NCBI] National Center for Biotechnology Information. 2008. NCBI Taxonomy.
(http://www.chem.missouri.edu/TannerGroup/people/white/NCBI%20taxonomy%20bro
wser/ecoli.htm).

Neidhardt, F. C., R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W.
S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger [eds.]. 1996. Escherichia
coli and Salmonellae: cellular and molecular biology, 2nd edition. ASM Press,
Washington, DC.

New, T. R. 1998. Invertebrate surveys for conservation. Oxford University Press, New York,
NY.

Nmorsi, O. P. G., G. Agbozele, and N. C. D. Ukwandu. 2007. Some aspects of epidemiology
of filth flies: Musca domestic, Musca domestic vicina, Drosophila melanogaster and
associated bacteria pathogens in Ekpoma, Nigeria. Vector Borne Zoonotic Dis. 7: 107-
117.

Ogden, I. D., N. F. Hepburn, and M. MacRae. 2001. The optimization of isolation media used
in immunomagnetic separation methods for the detection of Escherichia coli 0157 in
foods. J. Appl. Microbiol. 91: 1-7.

Olsen, A. R. 1998. Regulatory action criteria for filth and other extraneous materials: III.
Review of flies and foodborne enteric disease. Regul. Toxicol. Pharmacol. 28: 199-211.









Olson, T. A. and R. G. Dahms. 1945. Control of housefly breeding in partly digested sewage
sludge. J. Econ. Entomol. 38: 602-604.

Omisakin, R., M. MacRae, I. D. Ogden, and N. J. C. Strachan. 2003. Concentration and
prevalence ofEscherichia coli 0157 in cattle feces at slaughter. Appl. Environ.
Microbiol. 69: 2444-2447.

Ooka, T., J. Terajima, M. Kusumoto, A. Iguchi, K. Kurokwa, Y. Ogura, M. Asadulghani,
K. Nakayama, K. Murase, M. Ohnishi, S. Iyoda, H. Watanabe, and T. Hayashi.
2009. Development of a multiplex PCR-based rapid typing method for
enterohemorrhagic Escherichia coli 0157 strains. J. Clin. Microbiol. 47: 2888-2894.

Oporto, B., J. I. Esteban, G. Aduriz R. A. Juste, and A. Hurtado. 2008. Escherichia coli
0157:H7 and non-0 157 Shiga toxin-producing E. coli in healthy cattle, sheep and swine
herds in Northern Spain. Zoonotic Publ. Health 55: 73-81.

Orth, D., K. Grif, M. P. Dierich, and R. Wurzner. 2007. Variability in tellurite resistance and
the ter gene cluster among Shiga toxin-producing Escherichia coli isolated from humans,
animals and food. Res. Microbiol. 158: 105-111.

Osek, J. 2001. Molecular characterisation of Shiga toxin-producing Escherichia coli 0157
strains isolated in Poland. Int. J. Food Microbiol. 70: 175-177.

Pao, S., D. Patel, A. Kalantari, J. P. Tritschler, S. Wildeus, and B. L. Sayre. 2005. Detection
of Salmonella strains and Escherichia coli 0157:H7 in feces of small ruminants and their
isolation with various media. Appl. Environ. Microbiol. 71: 2158-2161.
Parker, R. R. 1916. Dispersion ofMusca domestic Linnaeus under city conditions in Montana.
J. Econ. Entomol. 9: 325-354.

Parkes, L. C. 1911. The common house fly (Musca domestica. J. Roy. Soc. Prom. Health 32: 4.

Paton, A. W. and J. C. Paton. 1998. Detection and characterization of Shiga toxigenic
Escherichia coli by using multiplex PCR assays for stx1, stx2, eaeA, enterohemorrhagic E.
coli hlyA, rJbo11i, and rfbo157. J. Clin. Microbiol. 36: 598-602.

Pearce, M. C., D. Fenlon, J. C. Low, A. W. Smith, H. I. Knight, J. Evans, G. Foster, B. A.
Synge, and G. J. Gunn. 2004. Distribution ofEscherichia coli 0157 in bovine fecal pats
and its impact on estimates of the prevalence of fecal shedding. Appl. Environ.
Microbiol. 70: 5737-5743.

Pedigo, L. P. and G. D. Buntin [eds.]. 1994. Handbook of sampling methods for arthropods in
agriculture. CRC Press, Boca Raton, FL.

Pell, A. N. 1997. Manure and microbes: public and animal health problems? J. Dairy Sci. 80:
2673-2681.









Pepper, H. J. 1944. Usefulness of microorganisms in studying dispersal of flies. Bull. U.S.
Army Med. Dept. 75: 121-122.

Perna, N. T., J. D. Glasner, V. Burland, and G. Plunkett III. 2002. The genomes of
Escherichia coli K-12 and pathogenic E. coli, pp. 3-53. In Donnenberg, M. S. [ed.].
Escherichia coli: Virulence mechanisms of a versatile pathogen. Academic Press, San
Diego, CA.

Perotti, M. A. and T. J. Lysyk. 2003. Novel growth media for rearing larval horn flies,
Haematobia irritans (Diptera: Muscidae). J. Med. Entomol. 40:22-29.

Petridis, M., M. Bagdasarian, M. K. Waldor, and E. Walker. 2006. Horizontal transfer of
Shiga toxin and antibiotic resistance genes among Escherichia coli strains in house fly
(Diptera: Muscidae) gut. J. Med. Entomol. 43: 288-295.

Pickens, L. G., N. O. Morgan, J. G. Hartsock, and J. W. Smith. 1967. Dispersal patterns and
populations of the house fly affected by sanitation and weather in rural Maryland. J.
Econ. Entomol. 60: 1250-1255.

Pickens L. G., N. O. Morgan, and R. W. Miller. 1972. Comparison of traps and other methods
for surveying density of populations of house flies in dairy barns. J. Econ. Entomol. 65:
144-145.
Pickens, L. G. 1989. Factors affecting the distance of scatter of house flies (Diptera: Muscidae)
from electrocuting traps. J. Econ. Entomol. 82: 149-151.

Pimentel, D. W. P. Nagel, and J. L. Madden. 1963. Space-time structure of the environment
and the survival of parasite-host systems. Am. Nat. 97: 141-167.

Pollack, K. 2005. Enhanced surveillance of haemolytic uraemic syndrome and other thrombotic
microangiopathies in Scotland, 2003-2004. Euro Surveill. 10:pii=2708.
(http://www.eurosurveillance.org/ViewArticle.aspx?Articleld=2708).

Porter, J., K. Mobbs, C. A. Hart, J. R. Saunders, R. W. Pickup, and C. Edwards. 1997.
Detection, distribution and probable fate of Escherichia coli 0157 from asymptomatic
cattle on a dairy farm. J. Appl. Microbiol. 83: 297-306.

Powell, D. 2008. The human face of E. coli 0157:H7: 3-year-old died in 2000.
(http://barfblog.foodsafety.ksu.edu/blog/138170/08/06/15/human-face-e-coli- 157H7-3-
year-old-died-2000).

Quarterman, K. D., W. Mathis, and J. W. Kilpatrick. 1954. Urban fly dispersal in the area of
Savannah, Georgia. J. Econ. Entomol. 47: 405-412.

Rasmussen, M. A., T. L. Wickman, W. C. Cray Jr., and T. A. Casey. 1999. Escherichia coli
0157:H7 and the rumen environment, pp. 39-49. In Stewart, C. S., and H. J. Flint [eds.].
Escherichia coli 0157 in farm animals. CABI, New York, NY.









Reisen, W. K. 2010. Landscape epidemiology of vector-borne diseases. Annu. Rev. Entomol.
55: 461-483.

Renter, D. G., S. L. Checkley, J. Campbell, and R. King. 2004a. Shiga toxin-producing
Escherichia coli in the feces of Alberta feedlot cattle. Can. J. Vet. Res. 68: 150-153.

Renter, D. G., J. M. Sargeant, L. L. Hungerford. 2004b. Distribution ofEscherichia coli
0157:H7 within and among cattle operations in pasture-based agricultural areas. Am. J.
Vet. Res. 65: 1367-1376.

Rice, D. H., K. M. McMenamin, L. C. Pritchett, D. D. Hancock, and T. E. Besser. 1999.
Genetic subtyping of Escherichia coli 0157 isolates from 41 Pacific Northwest USA
cattle farms. Epidemiol. Infect. 122: 479-484.

Rice, D. H., H. Q. Sheng, S. A. Wynia, and C. J. Hovde. 2003. Rectoanal mucosal swab
culture is more sensitive than fecal culture and distinguishes Escherichia coli 0157:H7-
colonized cattle and those transiently shedding the same organism. J. Clin. Microbiol. 41:
4924-4929.

Riemann, J. G., D. J. Moen, and B. J. Thorson. 1967. Female monogamy and its control in
houseflies. Insect Physiol. 13: 407-408.

Riley L.W., R. S. Remis, S. D. Helgerson, H. B. McGee, J. G. Wells, B. R. Davis, R. J.
Hebert, E. S. Olcott, L. M. Johnson, N. T. Hargrett, P. A. Blake, and M. L. Cohen.
1983. Hemorrhagic colitis associated with a rare Escherichia coli serotype. N. Engl. J.
Med. 308: 681-685.

Rodriguez, J. G. 1966. Axenic Arthropoda: current status of research and future possibilities.
Ann. New York Acad. Sci. 139: 53-64.

Rohde, M. 2008. Deal reached in E. coli death. Milwaukee Wisconsin Journal Sentinal Online.
(http://www.jsonline.com/news/milwaukee/29482674.html.

Rosef, 0. and G. Kapperud. 1983. House flies (Musca domestic) as possible vectors of
Campylobacterfetus subsp.jejuni. Appl. Environ. Microbiol. 45: 381-383.

Ruiu, L., A. Satta, and I. Floris. 2007. Susceptibility of the house fly pupal parasitoid
Muscidifurax raptor (Hymenoptera: Pteromalidae) to the entomopathogenic bacteria
Bacillus thuringiensis and Brevibacillus laterosporus. Biol. Control 43: 188-194.

Rutz, D. A., C. J. Geden, C. W. Pitts. 1994. Pest management recommendations for dairy
cattle. Cornell University and Penn. State Coop. Ext., Ithaca, NY.

SaccA, G. 1958. Research on speciation in the housefly. VI. Natural hybridism & experimental
hybridism between subspecies of Musca domestic L. Rend. Ist. Sup. Sanit. 21: 1170-
1184.









SaccA, G. 1964. Comparative bionomics in the genus Musca. Annu. Rev. Entomol. 9: 341-358.

SaccA, G. and L. Rivosecchi. 1958. Research on speciation in the housefly. V. Geographic
distribution of the subspecies of Musca domestic L. (Diptera: Muscidae). Rend. Ist. Sup.
Sanit. 21: 1149-1189.

Sanderson, M.W., J. M. Gay, D. D. Hancock, C. C. Gay, L. K. Fox, and T. E. Besser. 1995.
Sensitivity of bacteriologic culture for detection of Escherichia coli 0157:H7 in bovine
feces. J. Clin. Microbiol. 33: 2616-2619.

Sanderson, M., J. M. Sargeant, X. Shi, T. G. Nagaraja, L. Zurek, and M. J. Alam. 2006.
Longitudinal emergence and distribution of Escherichia coli 0157 genotypes in a beef
feedlot. App. Environ. Microbiol. 72: 7614-7619.

Sapers, G. M. and M. P. Doyle. 2009. Scope of the produce contamination problem., pp. 3-19.
In Solomon, E. B., and K. R. Matthews [eds.], The produce contamination problem:
causes and solutions. Academic Press, Elsevier Inc., Burlington, MA.

Sargeant, J. M., M. W. Sanderson, R. A. Smith, and D. D. Griffin. 2003. Escherichia coli
0157 in feedlot cattle feces and water in four major feeder-cattle states in the USA. Prev.
Vet. Med. 61: 127-135.

Sargeant, J. M., M. W. Sanderson, D. D. Griffin, and R. A. Smith. 2005. Factors associated
with the presence of Escherichia coli 0157 in feedlot-cattle water and feed in the
Midwestern USA. Prev. Vet. Med. 66: 207-237.

Sasaki, T., M. Kobayashi, and N. Agui. 2000. Epidemiological potential of excretion and
regurgitation by Musca domestic (Diptera: Muscidae) in the dissemination of
Escherichia coli 0157: H7 to food. J. Med. Entomol. 37: 945-949.

SAS Institute. 2002. SAS/STAT User's manual, Ver. 9.1. SAS Institute, Cary, NC.

Scaife, H. R., D. Cowan, J. Finney, S. F. Kinghorn-Perry, and B. Crook. 2006. Wild rabbits
(Oryctolagus cuniculus) as potential carriers of verocytotoxin-producing Escherichia
coli. Vet. Rec. 159: 175-178.

Schmidtmann, E. T., and P. A. W. Martin. 1992. Relationship between selected bacteria and
the growth of immature house flies, Musca domestic, in an axenic test system. J. Med.
Entomol. 29: 223-235.

Schoof, H. F. 1959. How far do flies fly, and what effect does flight pattern have on their
control: Pest Control 27: 16-18, 20, 22, 66.

Schoof, H. F., G. A. Mail, and E. P. Savage. 1954. Fly production sources in urban
communities. J. Econ. Entomol. 47: 245-254.









Schoof, H. F. and R. F. Siverly. 1954a. Multiple release studies on the dispersion of Musca
domestic at Phoenix, Arizona. J. Econ. Entomol. 47: 830-838.

Schoof, H. F. and R. F. Siverly. 1954b. Urban fly dispersion studies with special reference to
movement pattern ofMusca domestic. Am. J. Trop. Med. Hyg. 3: 539-547.

Schurrer, J. A., S. A. Dee, R. D. Moon, J. Deen, and C. Pijoan. 2006. Evaluation of three
strategies for insect control on a commercial swine farm. J. Swine Health Prod. 14: 76-
81.

Scudder, H. L. 1947. A new technique for sampling the density of house fly (Musca domestic)
populations. Publ. Health Rep. 62: 609-623.

Scudder, H. L. 1949. Some principles of fly control for the sanitarian. Am. J. Trop. Med. Hyg.
29: 609-623.

Sezonov, G., D. Joseleau-Petit, and R. D'Ari. 2007. Escherichia coli physiology in Luria-
Bertani broth. J. Bacteriol. 189: 8746-8749.

Sheppard, C. 1983. House fly and lesser fly control utilizing the black soldier fly in manure
management systems for caged laying hens. Environ. Entomol. 12: 1439-1442.

Shere, J. A., K. J. Bartlett, and C. W. Kaspar. 1998. Longitudinal study of Escherichia coli
0157:H7 dissemination on four dairy farms in Wisconsin. Appl. Environ. Microbiol. 64:
1390-1399.

Shura-Bura, B. L., A. D. Shaykov, YE. V. Ivanova, A. YA. Glazunova, M. S. Mitryukova,
and K. G. Fedorova. 1956. The migration of synanthropic flies to a town from the open.
Meditc. parzitolog. i parazit. boleznc. 4: 368-372.

Shura-Bura, B. L., O. I. Sukhomlinova, and B. I. Isarova. 1962. Use of radioactive tracers as
an aid to studying the ability of synanthropic flies to fly over water obstacles. Ft. Belvoir:

Skoda, S. R., G. D. Thomas, and J. B. Campbell. 1993. Abundance of immature stages of the
house fly (Diptera: Muscidae) from five areas in beef cattle feedlot pens. J. Econ.
Entomol. 86: 455-461.

Slutsker, L., A. A. Ries, K. Maloney, J. G. Wells, K. D. Greene, and P. M. Griffin. 1998. A
nationwide case-control study of Escherichia coli 0157:H7 infection in the United States.
J. Infect. Dis. 177:962-966.

Sokal, R. R. and R. L. Sullivan. 1963. Competition between mutant and wild-type housefly
strains at varying densities. Ecol. 44: 314-322.

Southwood, T. R. E. 1966. Ecological methods. Methuen and Co., London, UK.









Stafford III, K. C. 2008. Fly management handbook: A guide to biology, dispersal, and
management of the house fly and related flies for farmers, municipalities, and public
health officials. The Conn. Ag. Exp. Sta, New Haven, CT., Bull. 1013, May 2008.

Stein, W. 1986. Dispersal of insects of public health importance, pp. 242-252. In W.
Danthanarayana [ed.], Insect Flight: Dispersal and Migration. Springer-Verlag, Berlin,
Heidelberg.

Steinhaus, E. A. 1940. The microbiology of insects: with special reference to the biologic
relationships between bacteria and insects. Bact. Rev. 4:17-57.

Steinhaus, E. A. 1946. Insect microbiology. New York, NY, Hafner Publishing Co.

Steinhoff, U. 2005. Who controls the crowd? New findings and old questions about the intestinal
microflora. Immunol. Lett. 99:12-16.

Steinkraus, D. C., C. J. Geden, and D. A. Rutz. 1993. Prevalence of Entomophthora muscae
(Cohn) Fresenius (Zygomycetes: Entomophthoraceae) in house flies (Diptera: Muscidae)
on dairy farms in New York, and induction of epizootics. Biol. Control. 5: 405-411.

Stoetzel, M. B. 1989. Common names of insects and related organisms. Entomol. Soc. of
America, College Park, MD.

Suda, K. J., B. L. Love, T. J. Gladney, and K. W. Garey. 2003. Health and economic
outcomes of hospitalized patients with Clostridium difficile-associated diarrhea. Abstr.
Intersci. Conf. Antimicrob. Agents Chemother. (43rd meeting, abstract no. K-734).

Sukontason, K., K. L. Sukontason, R. C. Vogtsberger, N. Boonchu, T. Chaiwong, and S.
Piangjai. 2003. Prestomal teeth of some flies of medical importance. Micron 34: 449-
452.

Sulaiman, S. M. Z. Othman, and A. H. Aziz. 2000. Isolations of enteric pathogens from
synanthropic flies trapped in downtown Kuala Lumpar. J. Vec. Ecol. 25: 90-93.

Swadener, C. 1994. Bacillus i/un i/gie//i\ (B.t.). J. Pest. Reform 14: Fall 1994.
http://www.safe2use.com/poisons-pesticides/pesticides/BtK/btk.htm.

Szalanski, A. L., C. B. Owens, T. McKay, and C. D. Steelman. 2004. Detection of
Campylobacter and Escherichia coli 0157:H7 from filth flies by polymerase chain
reaction. Med. Vet. Entomol. 18: 241-246.

Taft, H. M. and H. R. Agee. 1962. A marking and recovery method for use in boll weevil
movement studies. J. Econ. Entomol. 55: 1018-1019.

Tarr, P. I. and M. A. Neill. 2001. Escherichia coli 0157:H7. Gastroenterol. Clin. Am. 30: 735-
751.









Taylor, C. E. and R. R. Sokal. 1976. Oscillations in housefly population sizes due to time lags.
Ecol. 57: 1060-1067.

Taylor, L. R. 1963. Analysis of the effect of temperature on insects in flight. J. Anim. Ecol. 32:
99-117.

Teplitski, M., J. D. Barak, and K. R. Schneider. 2009. Human enteric pathogens in produce:
un-answered ecological questions with direct implications for food safety. Curr. Op.
Biotechnol. 20: 166-171.

Tesch, M. J. and W. G. Goodman. 1995. Dissemination of microbial contaminants from house
flies electrocuted by five insect light traps. Inter. J. Environ. Health Res. 5: 303-309.

Thimijan, R. W., L. G. Pickens, N. O. Morgan, and R. W. Miller. 1972. House fly capture as
a function of number of traps in a dairy barn. J. Econ. Entomol. 65: 876-877.

Thomas, G. D.1993. The influence of beef cattle feedlots on the urban fly problem pp. 1-16. In
Thomas, G. D. and S. R. Skoda [eds.]. Rural flies in the urban environment? North
Central Regional Res. Bull. 335, Institute of Agriculture and Natural Resources Res.
Bull. 317. University of Nebraska Agricultural Research Division, Lincoln, NE.

Thomas, G. D. and S. R. Skoda. [eds.]. 1993. Rural flies in the urban environment? Research
Bulletin 317. North Central Regional Research Publication No. 335.

Thompson, F. C. 2009. Nearctic Diptera: twenty years later, pp. 3-46. In T. Pape, D. Bickel, and
R. Meier [eds.]. Diptera diversity: status, challenges and tools. Koninklijke Brill, Leiden,
The Netherlands.

Thorpe, C. M., J. M. Ritchie, and D. W. K. Acheson. 2002. Enterohemorrhagic and other
Shiga toxin-producing Escherichia coli. pp. 119-154. In Donnenberg, M. S. [ed.].
Escherichia coli: Virulence mechanisms of a versatile pathogen. Academic Press, San
Diego, CA.

Tobin, P. C. and C. W. Pitts. 1999. Dispersal of Muscidifurax raptorellus Kogan and Legner
(Hymenoptera: Pteromalidae) in a high-rise poultry facility. Biol. Control 16: 68-72.

Todar, K. 2008. Pathogenic E. coli. In Online textbook of bacteriology. Univ. Wisconsin-
Madison Dept. Bacteriol. (http://www.textbookofbacteriology.net/e.coli.html).

Tsuda, Y., H. Toshihiko, Y. Higa, K. Hoshino, S. Kasai, T. Tomita, H. Kurahashi, and M.
Kobayashi. 2009. Dispersal of a blow fly, Calliphora nigribarbis, in relation to the
dissemination of highly pathogenic avian influenza virus. Jpn. J. Infect. Dis. 62: 294-297.

Turchin, P. 1998. Quantitative analysis of movement. Sinauer Associates, Inc. Publ.
Sunderland, MA.









Turchin, P., F. J. Odendaal, and M. D. Rausher. 1991. Quantifying insect movement in the
field. Environ. Entomol. 20: 955-963.

Turner Jr., E. C. and P. L. Ruszler. 1989. A quick and simple quantitative method to monitor
house fly populations in caged layer houses. Poult. Sci. 68: 833-835.

Tutenel, A. V., D. Pierard, D. Vandekerchove, J. Van Hoof, and L. De Zutter. 2003.
Sensitivity of methods for the isolation of Escherichia coli 0157 from naturally infected
bovine faeces. Vet. Microbiol. 94: 341-346.

Tyler, H. L. and E. W. Triplett. 2008. Plants as a habitat for beneficial and/or human
pathogenic bacteria. Annu. Rev. Phytopathol. 46: 53-71.

[UF-EHS] University of Florida, Environmental Health and Safety. 2008. Biological safety.
(http://www.ehs.ufl.edu/bio/).

Unc, A., and M. J. Goss. 2006. Culturable Escherichia coli in soil mixed with two types of
manure. Soil Sci. Soc. Am. J. 70: 763-769.

[USDA:APHIS:VS] United States Department of Agriculture, Animal and Plant Health
Inspection Service, Veterinary Service. 1997. An update: Escherichia coli 0157:H7 in
humans and cattle. USDA:APHIS:VS Centers for Epidemiology and Animal Health. Fort
Collins, CO. (http://www.aphis.usda.gov/animalhealth/emergingissues
/downloads/ecoupdat.pdf).

[USDA-FSIS] Food Safety and Inspection Service, Office of Public Health Science. 2008a.
Detection, isolation and identification of Escherichia coli 0157:H7 from meat products.
Microbiol. Lab. Guide (MLG) 5.04. (www.fsis.usda.gov/PDF/MLG 5_04.pdf).

[USDA-FSIS] Food Safety and Inspection Service, Office of Public Health Science. 2008b.
Procedure for the use of Escherichia coli 0157:H7 screening tests. Microbiol. Lab. Guide
(MLG) 5A.01. (www.fsis.usda.gov/PDF/Mlg_5A_01.pdf).

Van Donkersgoed, J., T. Graham, and V. Gannon. 1999. The prevalence ofverotoxins,
Escherichia coli 0157:H7, and Salmonella in the feces and rumen of cattle at processing.
Can. Vet. J. 40: 332-338.

Van Donkersgoed, J., D. Hancock, D. Rogan, and A. A. Potter. 2005. Escherichia coli
0157:H7 vaccine field trial in 9 feedlots in Alberta and Saskatchewan. Can. Vet. J. 46:
724-728.

van Zwet, A. A., J. C. Thijs, A. M. D. Kooistra-Smid, J. Schirm, and J. A. M. Snijder. 1994.
Use of PCR with feces for detection of Helicobacterpylori infections in patients. J. Clin.
Microbiol. 32: 1346-1348.

Vidovic, S. and D. R. Korber. 2006. Prevalence of Escherichia coli 0157 in Saskatchewan









cattle: Characterization of isolates by using random amplified polymorphic DNA PCR,
antibiotic resistance profiles, and pathogenicity determinants. Appl. Environ. Microbiol.
72: 4347-4355.

Visetsripong, A., K. Pattaragulwanit, J. Thaniyavarn, R. Matsuura, A. Kuroda, and 0.
Sutheinkul. 2007. Detection of Escherichia coli 0157:H7 vt and rfbo17 by multiplex
polymerase chain reaction. Southeast Asian J. Trop. Med. Public Health 38: 82-90.

Void, L., A. Holck, Y. Wasteson, and H. Nissen. 2000. High levels of background flora
inhibits growth of Escherichia coli 0157:H7 in ground beef. Int. J. Food Microbiol. 56:
219-225.

Wallace, J. S. and K. Jones 1996. The use of selective and differential agars in the isolation of
Escherichia coli 0157 from dairy herds. J. Appl. Bacteriol. 81: 663-668.

Wang, L. and P. R. Reeves. 1998. Organization of Escherichia coli 0157 O antigen gene
cluster and identification of its specific genes. Infect. Immun. 66: 3545-3551.

Watanabe, Y., K. Ozasa, J. H. Mermin, P. M. Griffin, K. Masuda, S. Imashuku, and T.
Sawada. 1999. Factory outbreak of Escherichia coli 0157:H7 infection in Japan. Emerg.
Infect. Dis. 5: 424-428.

Watson, D. W., C. J. Geden, S. J. Long, and D. A. Rutz. 1995. Efficacy ofBeuveria bassiana
for controlling the house fly and stable fly (Diptera: Muscidae) Biol. Control 5: 405-411.

Watson, D. W., E. L. Nino, K. Rochon, S. Denning, L. Smith, and J. S. Guy. 2007.
Experimental evaluation ofMusca domestic (Diptera: Muscidae) as a vector of
Newcastle disease virus. J. Med. Entomol. 44: 666-671.

Watson, D. W. and J. J. Petersen. 1993. Seasonal activity of Entomophthora muscae,
(Zygomycetes: Entomophthorales) inMusca domestic L., (Diptera: Muscidae) with
reference to temperature and relative humidity. Biol. Control. 3: 22-26.

Watson, D. W., P.A.W. Martin, and E. T. Schmidtmann. 1993. Egg yolk and bacteria growth
medium for Musca domestic (Diptera: Muscidae). J. Med. Entomol. 30: 820-823.

Weather Underground. 2009. Daily summary history for KFLLAKEC8.
(http://www.wunderground.com/weatherstation/WXDailyHistory.asp?ID=KFLLAKEC8).

Wellington, W. G. 1945. Conditions governing the distribution of insects in the free atmosphere.
Can. Ent. 77: 7-15.

West, L. 1951. The housefly. its natural history, medical importance and control. Comstock
Pub., Ithaca, NY.









Wetzel, A. N. and J. T. LeJeune. 2006. Clonal dissemination of Escherichia coli 0157:H7
subtypes among dairy farms in Northeast Ohio. Appl. Environ. Microbiol. 72: 2621-
2626.

Wilde, J., J. Eiden, and R. Yolken. 1990. Removal of inhibitory substances from human fecal
specimens for detection of group A rotaviruses by reverse transcriptase and polymerase
chain reactions. J. Clin. Microbiol. 28: 1300-1307.

Wilkes, A., G. E. Bucher, J. W. M. Cameron, and A. S. West Jr. 1948. Studies on the
housefly (Musca domestic L.) I. The biology and large scale production of laboratory
populations. Can J. Res. Sec. 26: 26-56.

Williams, D. F. 1973. Sticky traps for sampling populations of Stomoxys calcitrans. J. Econ.
Entomol. 66: 1279-1280.

Williams, J. R. P. 1973. Dispersal of 32P-labelled adult Fannia canicularis. Int. Pest Cont. 15:
20-22.

Williams, D. F., C. S. Lofgren, and R. K. Vander Meer. 1990. Fly pupae as attractant carriers
for toxic baits for red imported fire ants (Hymenoptera: Formicidae). J. Econ. Entomol.
83: 67-73.

Willshaw, G. A., J. Thirlwell, A. P. Jones, S. Parry, R. L. Salmon, and M. Hickey. 1994.
Vero cytotoxin-producing Escherichia coli 0157 in beefburgers linked to an outbreak of
diarrhoea, haemorrhagic colitis and haemolytic uraemic syndrome in Britain. Lett. Appl.
Microbiol. 19: 304-307.

Wilton, D. P. 1963. Dog excrement as a factor in community fly problems. Proc. Hawaiian
Entomol. Soc. 18: 311-317.

Winpisinger, K. A., A. K. Ferketich, R. L. Berry, and M. L. Moeschberger. 2005. Spread of
Musca domestic (Diptera: Muscidae), from two caged layer facilities to neighboring
residences in rural Ohio. J. Med. Entomol. 42: 732-738.

Wong, T. T. Y. and M. L. Cleveland. 1970. Flourescent powder for marking deciduous fruit
moths for studies of dispersal. J. Econ. Entomol. 63: 338-339.

Xavier, B. M. and J. B. Russell. 2006. Bacterial competition between a bacteriocin-producing
and a bacteriocin-negative strain of Streptococcus bovis in batch and continuous culture.
FEMS Microbiol. Ecol. 58: 317-322.

Yates, W. W., A. W. Lindquist and J. S. Butts. 1952. Further studies of dispersion of flies
tagged with radioactive phosphoric acid. J. Econ. Entomol. 45: 547-548.









Yoruk, R., J. A. Hogsette, R. S. Rolle, and M. R. Marshall. 2003. Apple polyphenol oxidase
inhibitor(s) from the common house fly (Musca domestic L.). J. Food Sci. 68: 1942-
1947.

Zarchi, A. A. K. and H. Vatani. 2009. A survey on species and prevalence rate of bacterial
agents isolated from cockroaches in three hospitals. Vector Borne Zoonotic Dis. 9: 197-
200.

Zarrin, M. Z., B. Vazirianzadeh, S. S. Solary, A. Z. Mahmoudabadi, and M. Rahdar. 2007.
Isolation of fungi from housefly (Musca domestic) in Ahwaz, Iran. Pak. J. Med. Sci. 23:
917-919.

Zhao, T., M. P. Doyle, J. Shere, and L. Garber. 1995. Prevalence of enterohemorrhagic
Escherichia coli 0157:H7 in a survey of dairy herds. Appl. Environ. Microbiol. 61:
1290-1293.

Zurek, L., C. Schal, and W. Watson. 2000. Diversity and contribution of the intestinal bacterial
community to the development of Musca domestic (Diptera: Muscidae) larvae. J. Med.
Entomol. 37: 924-928.

Zurek, L., S. S. Denning, C. Schal, and D. W. Watson. 2001. Vector competence of Musca
domestic (Diptera: Muscidae) for Yersiniapseudotuberculosis. J. Med. Entomol. 38:
333-335.









BIOGRAPHICAL SKETCH

Roxanne Burrus is an active-duty medical entomologist in the United States Navy. After

completing her Ph.D. at the University of Florida, she will be reporting to Lima, Peru. Previous

duty stations have included Bangor, WA and San Diego, CA.

Roxanne's family has a history of military service. Both parents, James D. Burrus and

Dawn Elaine (Marcet) Burrus, served in the Navy, and her maternal grandfather, Henry Marcet,

served in the Army. She has one sister, Karen Elizabeth Burrus. Roxanne and her family have

lived in many different countries, including the United States, Taiwan, Cyprus, and Spain.

Roxanne has two bachelors' degrees; the first one in mathematics with a minor in computer

science from the University of Southern Mississippi (1988) and the second in biology with a pre-

medical emphasis from the University of Massachusetts at Amherst (2002). She completed a

master's degree in medical and urban entomology at the University of Florida (2004).

In spare time, which she had very little of while fulfilling the requirements of this Ph.D.

during the last three years, Roxanne likes to participate in triathlons and half-marathons. She was

a member of the UF Triathlon team, the Trigators, during both degree programs at the University

of Florida.





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1 MUSCA DOMESTICA L. (DIPTERA: MUSCIDAE) DISPERSAL FROM AND ESCHERICHIA COLI O157:H7 PREVALENCE ON DAIRIES IN NORTHCENTRAL FLORIDA By ROXANNE GRACE BURRUS A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2010

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2 2010 R oxanne G. Burrus

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3 To f amily and f riends

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4 ACKNOWLEDGMENTS Thanks to the following individuals who made completion of this dissertation possible Thanks to m y committee members Jerome A. Hogsette, Phillip E. Kaufman, James E. Maruniak, Volker Mai, and Amarat Simonne. Their const ructive suggestions provided insight and were tremendously beneficial at many levels Thanks to Heather Furlong, Lois Wood, Chris Geden, and Melissa Doyle for assistance with the fly dispersal study and house fly rearing. Thanks to Richard Robbins Marianne Radziewicz and David Hill of the Armed Forces Pest Management Board (AFPMB), for assistance with the literature Thanks to Wei Yea Hsu Luis Mendoza, James Becnel, and Julia Pridgeon, for assistance with microbiological processing of samples. Thanks to Dongyan Wang for statistical assistance Thanks to J ames E. Maruniak and Alejandra Garcia Maru niak, Michael Scharf, Volker Mai and Maria for assistance with polymerase chain reaction assays. Thanks to Debbie Hall, Katherine Smitherman and Mrs. Quispe for "over the top" administrative support. Thanks t o Navy Entomology personnel for mentor ing and guidance, including Stanton Cope, Gregory Beavers, George Schoeler David Hoel ; and from the Air Force, Doug Burkett On a more personal note, the following family members and friends provided timely support, advice, encouragement and inspiration : Dawn Bur rus, Ka ren Burrus, Kimmer O'Neill, Renee Edge, Christina Overstreet, Bonnie and Gene Parrish, Ulrich Bernier, Reginald Coler, and Eugene Gerberg Finally, thanks to all the persons involved with the US Navy's Duty Under Instruction (DUINS) program that pro vided the opportunity to attend graduate school while serving as a Naval preventive medicine officer.

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5 TABLE OF CONTENTS page ACKNOWLEDGMENTS ...............................................................................................................4 LIST OF TABLES ...........................................................................................................................8 LIST OF FIGURES .........................................................................................................................9 LIST OF ABBREVIATIONS ........................................................................................................10 ABSTRACT ...................................................................................................................................12 CHAPTER 1 INTRODUCTION AND LITERATURE REVIEW ..............................................................14 Introduction .............................................................................................................................14 True Flies ................................................................................................................................15 Medically and Economically Important Flies .........................................................................16 House Fly ................................................................................................................................16 Nomenclature ..................................................................................................................17 Origin and Distribution ....................................................................................................19 Classification and Taxonomy ..........................................................................................19 Biology and Ecology .......................................................................................................20 Life cycle .................................................................................................................21 Breeding substrates ..................................................................................................26 Nutrition and diet .....................................................................................................27 Bacteria as Food .......................................................................................................28 Microbiology ...................................................................................................................30 Microorganism persistence, replication and genetic transfer ...................................31 Development of antibiotic resistance in house fly gut .............................................33 Inse ct Movement .............................................................................................................35 Mark ReleaseRecapture Studies and Techniques ..........................................................36 Flight and Dispersal .........................................................................................................37 Mark and Release Techniques .........................................................................................39 Population Dynamics: Monitoring House Fly Populations .............................................41 Passive monitori ng methods .....................................................................................41 Active monitoring methods ......................................................................................44 House Fly Economical Impacts and Disease Outbreaks .................................................47 House Fly Management ...................................................................................................48 Cultural control ........................................................................................................48 Mechanical control ...................................................................................................49 Biological control .....................................................................................................51 Chemical control ......................................................................................................53 Integrated Pest Management ...........................................................................................54 Usefulness of House Flies ...............................................................................................55

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6 Enterobacteriaceae ..................................................................................................................57 Escherichia coli ......................................................................................................................58 Pathogenic Escherichia coli ...................................................................................................61 Escherichia coli O157:H7 ......................................................................................................63 Escherichia coli O1 57:H7 and Cattle ..............................................................................63 Escherichia coli O157:H7 Outbreaks ..............................................................................65 Escherichia coli O157:H7 Pathogenicity ........................................................................67 Escherichia coli O157:H7 Prevalence and Persistence ...................................................68 Escherichia coli O157:H7 Detection, Isolation and Identification .................................69 Escherichia coli O157:H7 and DNA based Isolation Techniques ..................................76 Summary .................................................................................................................................77 2 HOUSE FLY DISPERSAL ....................................................................................................78 Introduction .............................................................................................................................78 Materials and Methods ...........................................................................................................83 Laboratory Facilities and Rearing ...................................................................................83 Description of Study Area ...............................................................................................84 House Fly Collection and Rearing ..................................................................................88 Transport of Adult Flies to the Field ...............................................................................92 Marking, Releasing and Recapturing Adult House Flies ................................................92 Effects of Flu orescent Dust on House Fly Adults ...........................................................96 Weather ............................................................................................................................98 Statistical Analysis ..................................................................................................................99 Results .....................................................................................................................................99 Discussion .............................................................................................................................102 3 ESCHERICHIA COLI O157:H7 PREVALENCE ................................................................131 Introduction ...........................................................................................................................131 Materials and Methods .........................................................................................................134 DNA Quantification ......................................................................................................151 16S rDNA PCR Analysis ..............................................................................................152 Statistical Analysis ................................................................................................................152 Results ...................................................................................................................................154 Fly Monitoring ...............................................................................................................154 Enumeration of Aerobic Bacteria and Escherichia coli O157:H7 ................................155 Escherichia coli O157:H7 Preval ence by Direct Culture ..............................................156 Escherichia coli O157:H7 Prevalence by Polymerase Chain Reaction ........................157 Discussion .............................................................................................................................158 4 OVERALL CONCLUSIONS ...............................................................................................179 Conclusions ...........................................................................................................................179 Background ....................................................................................................................179 Conclusions ...................................................................................................................180 Future Research ....................................................................................................................181

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7 Summary ...............................................................................................................................181 LIST OF REFERENCES .............................................................................................................183 BIOGRAPHICAL SKETCH .......................................................................................................215

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8 LIST OF TABLES Table page 21 Mean and maximum distances flown per week and year by marked house flies released at a dairy in north central Florida. ......................................................................125 22 Trap distances (km) from the release site, the t otal numbers of marked house flies captured per alsynite sticky trap, and the cumulative percentage of marked flies.. ........126 23 Weekly recapture rate of marked house flies on alsynite sticky traps ............................128 24 Total number of house flies and marked house flies captured on alsynite sticky traps following release at a dairy farm in north central Florida. ............................................129 25 House fly release week and recapture rate and associated weekly weather data. ............130 31 Dates and house fly monitoring methods used at two Florida dairies. ............................173 32 Primer nucleotide sequences used to amplify target genes in PCR assay. ......................174 33 Enumeration of aerobic bacteria (CFU/g) using Petrifilm Aerobic Plate Count plates inoculated with 1 ml of the unenriched sample. ..............................................................175 34 Mean enumeration, by site and by sample type, of aerobic bacteria (CFU/g) using Petrifilm Aero bic Plate Count plates .. .............................................................................176 35 Prevalence (%) and number of E. coli O157:H7 CHROMAgar positive samples and number of CT SMAC and PCRpositive samples. ..........................................................177

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9 L IST OF FIGURES Figure page 21 Alsynite trap (Olson Products Inc., Medina, OH) placed at dairies and used to recapture on dairy and dispersing house flies. .................................................................122 22 Alsynite trap (Olson Products Inc., Medina, OH) locations and distance (km) from release point ....................................................................................................................123 23 Examples of dairycollected collected hous e flies following excessive treatment with two dusts to determine 24 h mortality effects. .. ..............................................................124 31 Scudder grid (45 x 45 cm) used to assess house fly populations on dairy farms. .........169 32 Spot cards at Dairy A's milk barn and at Dairy B .........................................................170 33 Spot and sticky cards. ....................................................................................................171 34 Sampling methods for each type of collected sample.. ....................................................172

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10 LIST OF ABBREVIATIONS ARS Agricultural Rese arch Service. Scientific research agency of the USDA BAMM FDA's Bacteriological Analytical Man ual BM Background microorganisms Refers to competing microorganisms that interfere with microbiological isolation of target organism BLAST Basic Local Alignment Search Tool. NCBI web based tool used to find similar regions between nucle otide or protein seq uences BMBL Biosafety in Microbiological and Biomedical Laboratories CDC Centers for Disease Control and Prevention. Agency of HHS CMAVE Center for Medical, Agricultural, and Veterinary Entomology, located in Gainesville, Florida. Research center of the US DA ARS CT SMAC Sorbitol MacConkey agar supplemented with ce fixime (15 g/l) and potassium tellurite (1.25 g/l). Selective agar used to isolate Escherichia coli O157:H7 from samples. Addition of antibiotics increases specificity over that of conventional S MAC DOC Department of Commerce EHS UF's Department of Environmental Health and Safety FDA United States Food and Drug Administration FMRU Flies and Mosquitoes Research Unit. R esearch unit at CMAVE GHFD Gainesville (larval) house fly diet. Standard rearing medium at the USDA ARS CMAVE for immature house flies. Consists of 50% wheat bran, 30% alfalfa meal, and 2 0% cracked corn (Hogsette 1992) HHS United States Departmen t of Health and Human Services IMS Immunomagnetic separation IO WH Institute for One World H ealth ITIS Integrat ed Taxonomic Information System MMWR CDC's Morbidity and Mortality Weekly Report NCBI NLM's National Center for Biotechnology Information NCDC NESDIS's National Climatic Data Center

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11 NESDIS NOAA's National Environmental Satellite, Data, and Information Service NI H National Institutes of Health NLM NIH's National Library of Medicine NOAA DOC's National Oceanic and Atmospheric Administration OHS CDC's Office of Health and Safety SMAC Sorbitol MacConkey agar Selective agar used to isolate Es cherichia coli O157:H7 from samples UF University of Florida USDA United States Department of Agriculture USN United States Navy WHO World Health Organization

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12 Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy MUSCA DOMESTICA L. (DIPTERA: MUSCIDAE) DISPERSAL FROM AND ESCHERICHIA COLI O157:H7 PREVALENCE ON DAIRIES IN NORTHCENTRAL FLORIDA By Roxanne Grace Burrus August 2010 Chair: Jerome A. Hogsette Cochair: Phillip E. Kaufman Major: Entomology and Nematology House fly, Musca domestica L., dispersal up to 3 km from a dairy release site into a nearby town was documented in this study. Dispersal occurred by both direct flig ht across multiple, mixed habitat types including open fields with interspersed tree copses, and by corridors and edges provided by local roads Marked flies were collected at an adjacent dairy, and at most traps that were set along two well travelled road s connecting the dairies and the town. Additionally, one marked fly was cap tured on a trap that was placed outside of a restaurant in town 3 km from the release site In total, 0.67% (250) of marked house flies were recaptured from a total release of 37,2000 marked flies over 11 wk. Escherichia coli O157:H7 was isolated from two dairies in north central Florida using immunomagnetic separation followed by direct culture plating onto CHROMAgar and sorbitol MacConkey agar supplemented with potassium tellurite and cefixime (CT SMAC) selective media. Presumptive identification of E. coli O157:H7 was confirmed by polymerase chain reaction (PCR) using the fliCH7 and rfbEO157 gene amplification. Dairy samples that were tested included pools of house flies, spilled grain, and fresh dairy cattle manure. Forty two percent (24/57) of samples were positive by direct culture using both

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13 CHROMAgar and CT SMAC agar, with 11 positive samples from CHROMAgar and 13 positive samples from CT SMAC agar The 24 positive samples were submitted to PCR; of those 24 samples, 14 (58%) were confirmed by PCR. Two of the PCR confirmed samples were falsenegatives on CHROMAgar media, but presumptive positive on CT SMAC media, indicating the importance of analyzing samples by more than one m ethod, and demonstrating the sensitivity of PCR. Direct culture prevalence rates from CHROMAgar were 1 4.0% (8/57) for house flies, 5.3% (3/57) for grain, and 0% (0/57) for manure. The rate of CHROMAgar i solation of E. coli O157:H7 from house flies was 2.6 times greater than from grain. The PCR confirmation rates were 67% (8 / 12) for house flies, 56 % (5/9 ) for grain and 33 % (1/3 ) for manure. These data suggest that detection of E. coli O157:H7 on dairies might be more accurately determined by testing house f lies instead of grain or manure samples, regardless of which isolation m ethod is utilized. Flies are easy to collect and process, and because they disperse into urban areas, they provide valuable information regarding a mobile element for pathogen transmis sion that is lacking in grain and manure samples. House flies should be an important consideration in the design of a pathogen monitoring program on dairies.

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14 CHAPTER 1 INTRODUCTION AND LITERATURE REVIEW Introduction With projected increases in global and local temperatures (West 1951, Borror et al. 1989, Gullan and Cranston 2000, Goulson et al. 2005, Meerburg et al. 2007) there will likely be a concomitant increase in insect vector populations, such as filth flies (Gratz 1999). This may potentially cause an increase in diarrheal diseases transmission by enteric bacteria such as Escherichi a coli Castellani and Chalmers, Shigella spp., and Salmonella spp., due to mechanical transmission of pathogenic organisms by filth flies (Greenberg 1971, Greenberg 1973). Filth fly life cycle developmental times for each developmental stage decrease in duration with i ncreased temperatures; as a result, the number of generations per year increases, and in temperate areas, can result in the establishment of year round fly populations. Higher thannormal, year round filth fly populations, living in close proximity to huma n populations, carrying viable antibiotic resistant bacteria in their digestive tracts, and are themselves resistant to multiple pesticides (Kaufman et al. 2001), present a tremendous potential for significant increase in human disease. Petridis et al. (20 06) documented E. coli O157:H7 genetic transfer of antibiotic resistance within the house fly, Musca domestica Linnaeus gut. Additionally, house flies can travel as far as 8 km (13 mi) from their breeding sites (Bishopp and Laake 1921, West 1951, Quarterm an et al. 1954, Sacc 1964, Stein 1986, Milio et al. 1988). This increases the potential for introduction of diseases across species, particularly from dairies with large house fly populations (Kaufman et al. 2005), to nearby human population centers. Due to anticipated expansi on of existing urban areas and the simultaneous expansion of animal facilities, large animal rearing facilities with concomitant filth fly populations will be increasingly in closer proximity to large human populations. The close proximity becomes a

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15 signif icant human health threat when the animal and human populations are within the filth flys normal flight distance, due to the increased opportunities available to the fly. This is of special concern in urban areas with hospitals producing biohazardous huma n waste, and in areas where both farms and urban centers generate antibiotic resistant bacterial pathogens which can then be transferred between sites. The goal of this research is to determine the public health threat posed by house fly, transmission of e nterohemorrhagic Escherichia coli across the rural urban interface between rural dairy farms and nearby urban residential areas. True Flies The house fly is a true fly. All true flies are in the Order Diptera, (Latin, di "two" + ptera "wings"), with hind wings that have been reduced to clubbed halteres. The halteres serve as balance sensory organs and project from the mesothorax at the site where hind wings would normally be located. Thus, true flies have only one pair of true wings (Borror et al. 1989). Possession of only one pair of wings, accompanied by the presence of halteres instead of the typical insectan characteristic of two pairs of wings, is a characteristic unique to Diptera, and serves as an easily observed visual diagnostic for differentiation of adult flies from most other adult insects. There are some exceptions to the onepair of wings characteristic: some insects in other orders, e.g., some mayflies (Ephemeroptera), have only one pair of wings, but lack halteres (Borror et al. 19 89). A mor e notable exception are male scale (Coccoideae) insects. Like Diptera, male scale insects possess halteres; however, scale halteres are bristled hooks instead of clubbed processes typical of true flies. Additionally, male scale insects lack mouthparts, and typically have one (rarely two) style like process projecting from the abdominal tip (Borror et al. 1989). There are 108 families of true flies (Olsen 1998), with an estimated 100,000 (Smith 1986) to 120,000 (Borror et al. 1989) identified species in the world. Identification of new species during the past 20 years has increased that number to more than 150,000 fly species (Thompson

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16 2009). Diptera are one of the largest orders of insects. Large, diverse populations are typical in most locations (Borror et al. 1989) with more than 20,000 species of Diptera in the Nearctic Region alone (Mullen and Durden 2002). Medically and Economically Important Flies Of the more than 150,000 true fly species (Thompson 2009), 350 (Greenberg 1971) are of medical, veterinary and/or economic importance to humans, either because they transmit the pathogens that cause disease, or because they interfere with animal rearing and crop production. Many of these flies are synanthropic, whereby they exhibit such strong associations wi th humans and domestic livestock that they are typically found living only with humans and in human environments (Borror et al. 1989). Fewer than 3.5% of flies representing fewer than 350 species in 29 families are associated with pathogen transmission. (G reenberg 1971). One species, M. domestica (Diptera: Muscidae), is so synanthropic that it is commonly referred to as the house fly (Howard 1900, Hewitt 1910, Hatch Jr. 1911, Parkes 1911, Mullen and Durden 2002) House Fly Flies have had a major impact during wars, due to transmission of enteric pathogens that cause diarrhea, dysentery and typhoid fever. Some war related examples follow. In 1898, the role of flies in pathogen transmission was discovered during the SpanishAmerican War (ANON 1940), as more soldiers died from typhoid fever, which is t ransmitted by house flies, than from battle injuries (Cirillo 2006). This discovery resulted in the application of the name typhoid fly to M. domestica and the publication of a 2 volume report by a commission of medical officers chaired by Walter Reed (A NON 1940). Combatants in the AngloBoer War, also waged in the late 1890s, suffered more deaths from typhoid fever than from battle injuries (Cirillo 2006). Subsequently, public health efforts were waged in the United States against the typhoid fly.

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17 Walte r Reed and his commission determined that M. domestica was predominantly responsible for the transmission of typhoid fever (ANON 1940, Cirillo 2006). During World War II, in 1942, United States Army camps suffered a massive epidemic of bacillary dysenter y. It was determined by serological and biochemical cultures that the same species of bacteria was present in the flies and in more than 91% of the infected soldiers. Importantly, no flies had been observed at the bivouac site prior to the Armys arrival. However, cases of dysentery and numbers of flies increased rapidly. After fly control measures were taken, the number of flies and the incidence of dysentery both decreased. Although no mortality occurred during this epidemic, 22% of the soldiers in the Division tested positive for the pathogenic organism. Infected persons were incapacitated for two to seven days. Changes in technology led to reduced breeding sites for house flies that in turn led to reduced numbers of flies and fewer concomitant cases o f diseases in which flies were implicated. For example, Graham Smith (1939), as cited in ANON (1940) attributed the decrease in summer diarrhea of infants to the increased use of automobiles with concomitant decreased use of horse drawn carriages. Nomencla ture Nomenclature of flies has varied over time, among countries, and between professional organizations. For example, in the United States, current convention for entomological terminology (Borror et al. 1989) in the largest North American entomological organization, the Entomological Society of America (ESA), specifies that common names for true flies include the word "fly" as a separate word: e.g., house fly, stable fly, horn fly, deer fly, horse fly, sand fly, etc. Correspondingly, this ESA convention specifies that insects in other orders, which are not true flies, append the suffix fly" to the common name root word: e.g., sawfly, stonefly, caddisfly, dragonfly, damselfly, etc. In contrast, older North American literature (before 1970),

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18 often cites the housefly," as one word (Matthysse 1945, Olson and Dahms 1945, Wilkes et al. 1948, West 1951, Floyd and Cook 1954, Sacc and Rivosecchi 1958, Sacc 1958, G reenberg 1959a ). Disparity of common names was a very big problem in much of the early literature, and can make historical research difficult. Two of the largest and most prominent entomological organizations in the United States formed standing committees just after the turn of the 20th century to resolve this matter: the American Association of Econo mic Entomology (AAEE) in 1903, and the American Entomological Society (predecessor to the current ESA) in 1907. The AAEE published the first list of common insect names in 1908, in which it formally established the use of "house fly," vs. "housefly," as the common name for M. domestica. In 1940, the AAEE and ESA insect nomenclature committees co published this list of authorized common insect names as a joint committee. In 1953, AAEE was absorbed into ESA and the resulting ESA has continued publication of t he list to the present. The list was originally published as a book (Stoetzel 1989, Bosik 1977) but is now available only online as database (ESA 2009). A more detailed description of the history of insect common name nomenclature is available for the int erested reader (Chapin 1989). Unfortunately, while adherence to the common names in this database is required for publication in ESA publications, many other peer reviewed publications do not comply with this attempt to unify nomenclature. This is particu larly true in medical publications, but is also true for some entomological journals that do not fall under ESA oversight. However, continued emphasis of standardized and unified common name nomenclature does seem to have succeeded in reducing the number o f publications using nonstandardized nomenclature; overall usage of

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19 insect common names is fairly well established throughout most of the ESA peer reviewed literature in the United States. In nonESA publications worldwide, "housefly" is still in use today (Zarrin et al. 2007, Malik et al. 2007, Black IV and Krafsur 2008, Cafarchia et al. 2009). Additionally, the reader will find multiple common names used for a particular insect. O ne will find that the same common name can refer to many different insects depending on local nicknames found in different geographical locations. There might be, as in the case of the house fly, multiple sub species of the insect that are so closely related they share not only their common name, but Linnaean classification as well. For example, Musca domestica vicina Macquart, sometimes referred to as M. vicina is the oriental house fly; however, it is often reported in the literature simply as "house fly," thus confusing it with M. domestica Origin and Distribution The hous e fly is a cosmopolitan pest that is strongly associated worldwide with human habitation, especially when domestic animals such as livestock, horses and poultry are present (West 1951). For all practical purposes, wherever humans live, the house fly also r esides. The house fly is excluded from arctic regions and higher altitudes due to extended cold temperatures (West 1951). Classification and Taxonomy Within the Order Diptera, the house fly is classified by Borror et al. (1989) as Suborder Brachycera, Inf raorder Muscomorpha (Cyclorrhapha), Division Schizophora, Section Calyptratae, Superfamily Muscoidea, Family Muscidae, Genus Musca and species domestica Linnaeus 1758. Each of these classification levels reflects some morphological trait that is character istic for M. domestica.

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20 The Suborder Brachycera contains the short horned flies (Borror et al. 1989), so named because they typically have short antennae with five or fewer segments. In contrast, flies the suborder Nematocera typically have long narrow antennae with 13 or more segments (Borror et al. 1989). Placement in Infraorder Muscomorpha (Cyclorrhapha) is based partially on the method of adult emergence from the puparium. The Muscomorpha includes the more advanced and specialized (higher) Diptera, which emerge through two slits that together appear T shaped on the anterior portion of the relatively stout, oval puparium (Jones 1977). Muscomorpha open these slits by alternately expanding and contracting a specialized inflatable sac (ptilinum) that is present on the head of emerging cyclorrhaphous flies. The pressure of the expanded ptilinum against the puparium causes the slits, only a few cells thick, to part so that a roughly circle shaped opening is formed from which the fly emerges. The ptilinum ret racts into the head shortly after emergence, but a seam (frontal suture) that is described both as being circular (Jones 1977), or as being shaped like a horse shoe (Hogsette, personal communication), remains visible in the adult fly (Jones 1977). The pres ence of a frontal suture in adult flies is responsible for classification into the Division Schizophora. In contrast, Aschiza adults typically lose their ptilinum and lack a frontal suture. Schizophora are divided into two sections (Calyptratae and Noncalyptratae) based on possession or lack of a pair of calypters (squamae, alulae) and a thoracic transverse suture (West 1951). Calyptrate flies possess well developed calypters that are large enough to conceal the halteres and they have an easily observed tra nsverse suture. Biology and Ecology The literature is replete with bionomic information concerning the house fly and closely related filth flies. Excellent monographs on M. domestica are available (Hewitt 1914, West 1951). A comprehensive monograph on the synanthropic black blow fly, Phormia regina Meigan, provides excellent information regarding the physiological aspects of eating that are

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21 also illustrative for M. domestica (Delthier 1976). Greenberg (1971, 1973) wrote a thorough twovolume series that sp ecifically addresses flies and pathogen transmission. Although much of the early literature is dated, these monographs provide tremendously useful information that is not easily available in the current literature. The wealth of information provided by the se authors is tremendous, and should not be neglected by anyone interested in studying the house fly or related muscids. Life cycle House flies have a holometabolous life cycle (Borror et al. 1989) which consists of four stages: egg, larva, pupa, and adult. Development of house flies at each life stage is temperaturedependent, so that the entire life cycle can be completed in as few as 8 d in the summer (West 1951). However, the typical life cycle for development from egg to adult in temperate zones ranges from 1014 d. First stage larvae hatch from the eggs within 812 h after oviposition. The three larval stages are typically completed as fol lows: first instar in 20 h 4 d, second instar in 24 h to several days, and third instar in 39 d. The pupal stage lasts an average of 5 d, although adverse conditions may extend this to several weeks. The adults live several weeks, with females generally living longer than males. During summer, adults may live 2 3 wk while cooler temperatures during spring and fall may contribute to adult longevity for up to 3 mo (West 1951) House fly eggs have a glossy white (West 1951) or creamy color (Mullen and Durden 2002), are approximately 1 mm in length (West 1951), but can range in size from 0.8 2.0 mm (Mullen and Durden), and are approximately 0.25 mm in width (West 1951). House fly eggs exhibit a slight concave indentation on the dorsal side that is due to the presence of two longitudinal ridges (ribs) that taper towards each other anteriorly (West 1951, Mullen and Durden 2002) to form hatching pleats (Mullen and Durden 2002). Gravid females disperse their

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22 eggs methodically throughout available breeding substra tes, depositing them either individually or in masses (West 1951). The females walk over and crawl carefully through the breeding substrate, seeking cracks and crevices into which they extend their ovipositor to dispense ova. The ova are laid so that they "rest on their broader posterior ends" (West 1951). Females oviposit 48 d after mating, and may take up to 24 h to deposit each batch of 100150 eggs. A total lifespan deposition of 46 batches of eggs can occur when flies are permitted to oviposit every two weeks (West 1951). Flies that are maintained in colony are induced to lay eggs every other day after they are five days of age (Hogsette, personal communication). House fly larvae are also called maggots ( Moon 2002). House fly larval length increases approximately 25 percent during each of three molts and body weight can increase 54fold in 4 d (West 1951). Third instars are approximately 12 mm (West 1951) to 15 mm (Moon and Meyer 1985) in length. House fly larvae possess two fused (Roback 1944) mouth hooks in their reduced (Moon 2002), unsclerotized head (Borror et al. 1989) that is located at the anterior, pointed end of a tapered body. The head lacks eyes but possesses small papillaelike antennae (Moon 2002). The larvae are creamy white in color (Moon 2002). The mouth hooks move in a vertical plane (Borror et al. 1989), providing physical evidence that these larvae are non predaceous, because their mouth hooks do not permit grasping. Instead, these mouth hooks permit the larvae to pull themselves th rough the media in which they live. House fly larvae are primarily sarcophagous, i.e., they feed on rotting organic matter (West 1951) (Gr. sapros rotten; phagein, to eat). House flies are also reported to be coprophagous, i.e., they feed on dung (Hammer 1941) (Gr. kopros dung). The paired mouth hooks can be retracted into the oral cavity or extended out, and are used to help the larva move through food substrates (Moon 2002). House fly larvae are legless (Borror et al. 1989). Each of the eight abdominal segments has a transverse row of spines on the ventral

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23 surface (creeping welts) that are used to facilitate movement (Moon 2002). House fly larvae possess a pair of lateral spiracles on the prothorax of second and third instars and a pair of spiracles at t he last abdominal segment of all instars (Moon 2002). The posterior (caudal) spiracles and associated structures are useful for aging and identifying muscid immatures, especially in third stage larvae (Moon 2002). Posterior spiracles are located on a spir acular plate which is encircled either completely or partially by a sclerotized peritreme. Developmental age can be determined to instar by counting the number of spiracular slits. First and second house fly instars possess two spiracular slits inside the spiracular plate, while third instars possess three slits. House fly larvae can be identified to species by examining the shape and orientation of the spiracles, the location of the molt scar, and the peritreme shape. House fly larvae have sinuous spirac les partially surrounding a large molt scar. The peritreme is completely sclerotized and encircles the spiracular area, with the peritreme flattened along the interior edge and the molt scar located mid way along the interior flattened edge of the peritrem e. Closely related muscid larvae such as the stable fly, Stomoxys calcitrans (L.), with similar body shapes and sizes can be differentiated by s shaped spiracular slits surrounding a small molt scar that is located in the center of a spiracular plate that lacks a flattened interior edge (Moon 2002). Mature 3rd instars migrate from the deeper moist, fermenting parts of the larval medium up into drier crusty toplayers (Greenberg 1959d). Once there, they wander for approximately 24 hours, during which time they cease feeding in preparation for pupation so that the prepupal gut is empty and contracted (Greenberg 1959d). The third instars integument becomes hardened during the process of pupariation, to form a puparium, in which the fly will complete its development (Moon 2002). House fly pupae

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24 are accurately called puparia (West 1951), to reflect the enclosure of the exarate (coarctate) pupa within the sclerotized last larval cuticle, in contrast to larval metamorphosis within a specially constructed pupal s tructure that is typical for insects other orders such as Lepidoptera and Coleoptera (Gullan and Cranston 2000). The puparium is red to brown in color and barrel shaped (Moon and Meyer 1985) and typically 6.3 mm in length (West 1951). Pupariation takes at least 6 h, with sclerotization gradually darkening the integument to its final dark brown color during this process (West 1951). The pupal stage typically lasts 5 d (West 1951), but can range from 310 d (Moon and Meyer 1985), depending on temperature (We st 1951, Moon and Meyer 1985). During adverse conditions, the pupal stage can last several weeks, and pupae may be able to hibernate (West 1951). Pupae are usually located in a cooler (West 1951) and drier (Moon and Meyer 1985) location than feeding larvae Third instar nonfeeding prepupae disperse to find suitable locations for pupal development. Large pupal aggregations numbering in the thousands can be found close to the medium surface and in surrounding soil (Moon and Meyer 1985). Pupal development las ting 4 d at 35 C (Greenberg 1959b) occurs with daily physiological changes. Within the first day, the pupa molts, and is encased in a transparent molting membrane with the head uneverted. On the second day, the head everts, and the compound eyes are amber On the third day, the eyes are orange or brown and bristles exhibit slight pigmentation. On the fourth day, the fly ecloses (Greenberg 1959b). House fly adults are small to medium sized flies approximately 6mm in length (West 1951). However, they may ra nge in size from 310 mm (Byrd and Castner 2009) or from 412 mm (Moon 2002). Numbers of adults and adult size are reduced by inadequate larval nutrition (West 1951) or by high larval population density (Sokal and Sullivan 1963, Black and Krafsur

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25 1985). De nsity induced adult size changes can influence mating success (Baldwin and Bryant 1981). The house fly life cycle length can vary due to one or a combination of multiple factors such as: temperature (Hewitt 1910), relative humidity (Hewitt 1910), and light (Greenberg 1959e). Additional factors that can influence developmental times include geographical region (Black and Krafsur 1968); season (LaBreque et al. 1972); breeding substrate composition (Skoda et al. 1993), pH, and moisture content (Evans 1916); population density ( Haupt and Busvine 1968, Taylor and Sokal 1977); and nutrition (West 1951). House flies begin their life cycle shortly after adult emergence. Males and females emerge in approximately equal proportions when adequate nutrition is availa ble (West 1951). When insufficient nourishment is available, significantly smaller males develop and these greatly outnumber the females (West 1951). The time required for sexual maturity in newly emerged adult house flies was reported as 1 d by Riemann e t al. (1967). Murvosh et al. (1964) found that males are sexually mature at 16 h and females at 24 h. A great deal of research has been conducted to determine the preoviposition period of house flies, for the purpose of concentrating control efforts agains t adults before oviposition occurs (Hutchison 1916). Hutchison (1916) determined that preoviposition periods range from 2.5 23 d, and are greatly influenced by temperature. His data imply that preoviposition periods also may be influenced by humidity and adult diet. Bishopp et al. (1915) also studied preoviposition periods, and they observed a seasonal / temperature influence on the preoviposition period, i.e. the period between emergence and oviposition, during which the fly becomes sexually mature and copulation takes place (Mellor 1919). Bishopp et al. (1915)

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26 observed preoviposition periods of 4 d in summer vs. more than 10 d in spring and fall, and Hewitt (1910) observed long preoviposition periods of up to 14 d. Several days elapse between sexual mat urity and copulation, so that females typically mate when 3 4 d old (West 1951). Adult female house flies are typically monogamous, with only a small percentage mating more than once, and none more than a few times. Monogamy appears to occur due to a thre shold limit for the receipt of seminal fluid (not of sperm) which can be reached by interrupted matings. Matings with castrated males can also result in loss of receptivity. Females may become sexually receptive again after 20 consecutive days of ovipositi on. Transfer of sperm is essentially completed within 10 minutes, although flies remain coupled for approximately 1 h. Even those adult female flies that mate more than once restrict their copulations to only a few times (Riemann et al. 1967) Flies copulate with the smaller male positioned directly above and clasping the female, with both insects facing the same direction, and with their abdominal t ips connected (West 1951). Breeding substrates House flies are cosmopolitan opportunists, and are associated with a wide variety of decomposing organic materials (Lole 2005). Availability of a natural substrate is the most important factor for house fly s election at any particular location and in a specific moment. Natural substrates consist of a tremendous variety of materials, including manure from cattle, horses, sheep, dogs, and humans; spilled grains; compost; garbage and landfills (Lole 2005) House flies are ubiquitously abundant on cattle farms, due to the availability of preferred breeding sites such as decaying grain based feed and manure (Skoda et al. 1993). Larvae spend their entire developmental lifecycle within the substrate selected by the gravid females. In general, house fly larval abundance in substrates reflects localized abundance of that particular substrate. F or example, prior to the development of the automobile, enormous

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27 house fly populations were observed in horse manure that was greatly abundant in urban streets. Post automobile, we find very few references to house flies breeding in urban horse manure. Cur rently, we find that house fly problems at animal rearing facilities, rather than in urban streets, dominate the literature. Current fly problems coincide with increased numbers of ever enlarging commercial facilities, which struggle to manage the copious amounts of fecal waste generated by the intensivelyreared animals. Nutrition and d iet Survival rate at each developmental stage, pupal weight, and adult fecundity are important measures of the nutritional status of reared flies. Larval populations that d o not receive adequate nutrition usually produce smaller flies with higher proportions of males. House fly larvae are successfully maintained in the laboratory on a variety of diets. After nutritional provision, important factors to consider in selecting a rearing medium are easeof use, expense, and inhibition of molds and fungi. Sukhapanth et al. (1961) compared fly development in a synthetic diet (100 g rice bran and husk, 350 g dry, low fat powdered milk, 75 icing sugar, 15 g Bakers yeast and 200 ml of 2% KOH in normal saline solution), a natural diet (fresh cow meat), and a combination of the two. House fly egg production and survivorship was highest at all developmental stages on the combined synthetic/meat diet and lowest at all stages on the sy nthetic diet. Sukhapanth et al. (1961) did not describe pupal weights, or adult sizes, so that comparison to diets used by other researchers cannot be made without repeating Sukhapanths work. Hogsette (1992) created the Gainesville house fly diet (GHFD) containing 30% alfalfa, 50% wheat bran, and 20% corn meal to take advantage of year round locallyavailable feed components. Mean larval weight (15 g) and adult eclosion rates from the GHFD diet did not differ significantly from that of the Chemical Specia lties Manufacturers Association larval

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28 medium (CSMA) (Greenberg 1959b). CSMA is composed of 33% wheat bran, 26.7% alfalfa meal, and 40.0% brewers grain (Greenberg 1959b). Additional benefits that Hogsette (1992) described for use of GHFD over CSMA were d ecreased costs, reduced delivery times, decreased storage time, increased feed quality, and suitability for stable fly, Stomoxys calcitrans L. larvae by addition of peanut hulls to the GHFD in equal volumes. Bacteria as Food The nutritional needs of house fly larvae are fulfilled by consumption of microorganisms and other substances located within the decomposing organic matter that also serves as a breeding substrate. Some natural components of substrates used for larval development of M. domestica are m icroorganisms, such as bacteria, viruses, and parasites. Postulation that house flies eat bacteria, and not just the decomposing substrate materials used for larval development, led to experiments to rear M. domestica in various synthetic media. Schmidtmann and Martin (1992) determined that house fly larvae depend either on the bacteria or bacterial metabolic products for essential nutrients. In some studies, agar based systems with known microbial organisms have been used to rear house fly larvae (Schmidtm ann and Martin 1992, Watson et al. 1993, Lysyk et al. 1999) and larvae were observed during development (Perotti and Lysyk 2003) Bacterial communities of decomposing organic substrates can change rapidly, due to a series of complex biotic and abiotic interactions (Archer and Young 1988, Jiang et al. 2002). Environmental conditions such as temperature and relative humidity affect several substrate factors, including decomposition rate, available water content, pH, and availability of oxygen. Bacterial com munities can also be impacted by the metabolic processes of the competing microorganisms (Jiang et al. 2002).

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29 Investigation into microbial contributions towards house fly larval development has led to experiments in which many specialized diets are used f or rearing the larvae. These specialized diets are narrowly defined by specific nutritional terminology such as "axenic," "gnotobiotic," "holidic, and "meridic. Because some researchers are describing the substrate as a rearing medium rather than definin g the nutritional composition of the medium, words such as "aseptic" and "sterile" have alternatively been used. Usage of these words is not consistent among researchers, and comparison of rearing methods must be carefully assessed by perusal of the scient ist's methods rather than by the word used to describe the larval rearing medium. Some subtle, but important differences separate each of these words. Although "axenic" and "aseptic" are frequently used synonymously in arthropod literature (Rodriguez 1966) ; both terms refer to the rearing of target organisms with no other living organisms present, e.g., rearing steri lized house fly in media containing only inert nutrients such as synthetic diets or nutrient agars. The composition of inert components might or might not be specified. Sterile is synonymous to axenic and aseptic, as seen in the rearing of house fly larv ae on synthetic egg yolk media and on blood agar plates that contained no other living organisms (Watson et al. 1993). Gnotobiotic refers to rearing of the target organism with only one other living organism present, e.g.: a gnotobiotically reared house fly is an aseptic fly that was fed known quantities of precisely defined foods, such as pure cultures of either Salmonella typhimurium or Proteus mirabilis in known concentrations (Greenberg et al. 1970, Watson et al. 1993). Many studies that are termed axenic would fall under the more specific term gnotobiotic due to provision of bacteria species; thus, it seems that the term gnotobiotic has been generally supplanted by the term axenic. Regardless of gnotobiotic status, axenic studies provide informati on about nutritional responses (Rodriguez 1966) and have been used to determine the relationship of

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30 bacteria and immature fly development by many researchers including Schmidtmann and Martin (1992), Watson et al. (1993), and Lysyk et al. (1999). The term s holidic and meridic were introduced by Dougherty (1959), as cited in Rodriguez (1966), to differentiate between diets that contain only components with completely known chemical structures (holidic) or diets that add components with unknown chemical structures, such as agar, to a holidic base (meridic) (Rodriguez 1966). Microbiology Musca domestica hosts a large variety of bacterial fauna, both internally and externally (Greenberg 1973). Due to the house flys association with a wide variety of fecal and decomposing organic materials, what can be considered natural fauna versus environmentally obtained fauna is unclear. Despite this, Greenberg (1959c) postulated that Proteus vulgaris, Proteus mirabilis Aerobacter aerogens and Escherichia freundii mig ht be considered the normal flora of house flies, after he isolated these bacteria from both laboratory flies reared on the CSMA diet and from natural populations of flies collected from horse manure. He further concluded that these bacteria are probabl y the predominant species in CSMA The advent of the microscope conferred the ability to study microorganisms, while culture methods provided the ability to grow microorganisms. Used together, these two technologies contributed greatly to the progress that has been made in isolating pathogens from M. domestica and related flies. Since then, the house fly has been determined to be capable of transmitting more than 100 microorganisms, including bacteria, parasites, viruses, and yeasts (Greenberg 1973). Many of the microorganisms isolated from house flies are pathogenic to humans.

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31 Microorganism p ersistence, r eplication and genetic t ransfer Extensive research has been conducted to examine the ability of microorganisms to survive (persist) (Ledingham 1911; Graha m Smith 1912; Greenberg 1959b, 1959c; Sasaki et al. 2000; Zurek et al. 2001; Nayduch et al. 2005) and multiply (replicate) within the house fly's alimentary canal and associated organs (Petridis et al. 2006, Macovei et al. 2008). The ultimate goal is to de termine the potential for biological (versus merely mechanical) transmission of pathogenic agents by the house fly. Microorganisms are consumed by larval house flies which depend on the pathogens or their metabolic products for growth and development ( Greenberg 1959b). Microorganism survival from the larval to the adult fly stages could result in increased dissemination by the house fly (Greenberg 1959b,d,e). Greenberg (1959c,d,e) observed large declines in the number of bacteria in the prepupae and emergent adults. He attributed the decline in prepupae to a cessation of feeding with continued excretion prior to pupation. This was confirmed by observation that the house fly prepupal gut is empty and contracted in contrast to the fully distended gut of ac tively feeding larvae (Greenberg 1959d). The decline of bacterial counts in newly emergent flies was attributed to the molting of foregut and hindgut during pupation (Greenberg 1965). This was confirmed by counting the number of bacteria present in shed puparia (105) versus the number of bacteria on the newly emerged flies (102). Greenberg (1965) reported that the bacteria survived the pupal period within the shed foregut and hindgut portions outside of the pupal fly, and were separated from the fly by a thin membrane. Therefore, adult flies emerged from the puparium with relatively few bacteria. Those bacteria that did remain within the fly were in the midgut, which is not shed during molting (Greenberg 1965). Bacteria were reported to survive in the adult fly for up to 12 h on the surface, and up to 30 h in the gut and feces (Grbel et al. 1997). Sasaki et al. (2000) reported survival of

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32 microorganisms for up to 3 d in the mouthparts and up to 4 d in the crop. The adult house fly mouthparts consist of a pr oboscis, which has two fleshy labella that can be spread out, sponge like, over food surfaces, to implement a suctorial action for uptake of liquids and minute food particles. Finally, the proboscis can be retracted into the head capsule, thus providing a micro habitat for bacteria that is relatively protected from desiccation and UV light (Kovacs et al. 1990). The proboscis also has one row of 56 three cusped (trifurcated) prestomal teeth on each side of a central food channel (Macloskie 1880, Broce and E lzinga 1984, Sukontason et al. 2003) These prestomal teeth scrape food resources, and are important for enhanced pathogen digestion (Kovacs et al. 190). Microorganism survival within the house fly gut seems to differ for different microorganisms. Nayduch et al. (2005) reported survival of Aeromonas caviae and Serratia liquefaciens in the alimentary tract for up to 5 d. Nayduch et al. (2005) described production of a baglike peritrophic membrane structure around feces (fecal pellet). They observed that these bacteria were located in the peritrophic membrane folds rather than in the peritrophic space, and not within the ectoperitrophic space; indicating that they were not transported out with the feces with the peritrophic membrane. They suggest that these bacteria are capable of evading entrapment by the peritrophic membrane by some method (Nayduch et al. 2005). Viable excretion of Yersinia ps uedotuberculosis was reported for up to 36 h with contamination of the environment detected for up to 30 h after inoculation by Zurek et al. (2001). Grbel et al. (1997), the first to detect viable Helicobacter pylori in house fly feces, reported survival of H. pylori for up to 12 h on the exoskeleton, 30 h in the gut, and 30 h in the feces. More than 100 spp. of bacteria have been isolated from adult house flies (Greenberg 1971), and from the adults alimentary tract (Ahmad and Zurek 2006). The number of bacteria

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33 on adults varies for different locations and bacteriological isolation methods. Sulaiman et al. (2000) isolated nine spp. of bacteria, including the first reported isolation of Burkholderia pseudomallei from house flies in Kuala Lumpar, Malaysia Successful replication of bacteria within the digestive tract (Macovei and Zurek 2007) and associated organs could play an important role in genetic transfer of virulence factors and antibiotic resistance from one bacterium to another, including transfer to diffe rent species, because bacteria often exchange genetic material during replication. This is particularly true of gram negative enteric bacteria such as E. coli sp., which possess extra chromosomal plasmids (Johnson 2002). Development of antibiotic r esistance in house fly gut Insect guts provide an ideal envir onment for exchange of genes by plasmid transfer and transconjugation between bacteria. The house fly has been implicated as a factor in the transfer of antibiotic resistance genes among bacteria (Macovei and Zurek 2006) because bacteria replication, which is usually accompanied by genetic exchange, occurs in the house fly gut (Petrides et al. 2006). Additionally, antibiotic resistance was observed on organic pig farms where antibiotics were not used, but fly numbers were large (Meerburg et al. 2007). Ther e is evidence that E. coli O157:H7 exchanges genetic material such as virulence factors and antibiotic resistance by plasmid transfer (Perna et al. 2002) or phagemediated transduction (Johnson 2002). Plasmid transfer of antibiotic resistance genes has led to disease outbreaks that were difficult to control because of development of multidrugresistant Staphylococcus aureus (MRSA) (Perna et al. 2002). This bacterial exchange of genetic material can occur within the house fly alimentary tract, which provides an environment conducive to the evolution and emergence of new pathogenic bacterial strains (Petrides et al. 2006). Exchange of antibiotic resistance between infectious bacteria is responsible for the worldwide increase in

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34 nosocomial (hospital acquired) i nfections that are resistant to all known antibiotics (Weinstein 1998). The potential for bacterial exchange of genetic material within the house fly gut presents potentially serious complications for any disease caused by pathogens that are disseminated b y house fly movement within a hospital environment. The potential for development of resistance to multiple antibiotics is particularly troublesome if the flies have access to the pathogens natural reservoirs; for example, access to dairy cattle for trans mission of E. coli O157:H7. The potential for flyborne bacteria dissemination coupled with bacterial exchange of antibiotic resistance is of concern in regards to the development of nosocomial infections (Boulesteix et al. 2005). In hospitals, intensive care units serve as the epicenter for these infections (Weinstein 1998). While hospitals and animal rearing facilities, such as dairies, both report increased antibiotic resistance due partially to over use (drugs to humans for treatment of bacterial infec tions and inclusion of drugs in animal feed for use as a growth stimulator), dispersal of flying insects such as the house fly between hospitals and dairies also could introduce new antibiotic resistance genes in each site. The potential for pathogen transmission, including antibiotic resistance, plus the possibility of economic losses due to infestations of synanthropic flies, has led to attempts to reduce fly populations wherever flies aggregate or build to annoying levels. Insects that harm human or ani mal health or damage valued resources are considered pests (Foster and Harris 1977). The importance of pest control for reduction of disease outbreaks is highlighted by reports of paired reduction of fly populations and a corresponding decrease in the dise ase incidence (Nash 1909, Lindsay and Scudder 1956, Cohen et al. 1991). This permits an estimated quantification of the house flys role in pathogen transmission. For example, Emerson (1999) reports a 75% reduction of muscid flies corresponding to a reduct ion in the number of new cases

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35 of trachoma and a 2226% reduction of muscid flies corresponding to a reduction in the number of new cases of diarrhea for villages in Gambia after fly control efforts reduced the numbers of muscid flies by approximately 75%. Insect Movement Terrestrial insect movement can occur by walking or by flying. Flight is used to both disperse and migrate (Angelo and Slansky 1984). Dispersal is typical of small, r strategist insects that produce large numbers of short lived offspring: movement out of a location with high population densities prevents overcrowding and decreases competition for resources. Dispersal is somewhat localized, so that insects disperse from one resource to another within a limited geographical range: for example, house flies and stable flies. Dispersal of insects may be similar to diffusion of small particles in air, so that dispersal may be randomized, and may even go back and forth between food/breeding resources during the insect's life span (Schoof and Sive rly 1954). In contrast, migration is typical of larger, longer lived K strategists that produce fewer offspring. Migration generally involves longer distances, is uni directional, and is a permanent relocation. In insects, migration usually occurs over mul tiple life stages and/or generations: for example, monarch butterflies. However, the differentiation between dispersal and migration can be blurred, as is seen for locusts which migrate over long distances, with an environmentally induced morphologically c hanged body, in response to limited food resources (Angelo and Slansky 1984). Although house flies are not thought to migrate, they do disperse frequently and readily. Adult house fly behavior and dispersal is influenced by climatic factors (also referred to as "environmental influences") such as temperature (Hewitt 1910), relative humidity (Hewitt 1910), barometric pressure, light intensity (Greenberg 1959b), and electrostatic fields (Johnson 1969). Dispersal of house flies and other synanthropic flies is of great interest, especially in livestock

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36 and public health situations, for both economic and medical reasons. Dispersal of house flies is frequently studied through markrelease recapture studies. Mark R eleaseR ecapture S tudies and T echniques Mark rele ase recapture (MRR) techniques are used to study insect dispersal, because they provide point to point dispersal information about insect movement from a release site to a recapture site. However, they do not describe the method or the path used (Turchin et al. 1991) MRR studies have been conducted using different substances to mark the flies. Ideally, a marker is not ubiquitous in the natural environment, but is quick and easy to apply, remains detectable for a long period, does not adversely affect fly behavior or mortality, is inexpensive, readily available, and has a long shelf life (Turchin 1991). Flies are marked using many different meth ods. In previous studies, flies have been made radioactive (by digestion of P 32 radiolabelled diets) and been individually handpainted, or dusted with fluorescent dusts (Wong and Cleveland 1970). Another marking technique involves amputating specific com binations of body parts and/or notching the exoskeleton in specific locations. Of these techniques, fluorescent dusts are the most convenient to use (Hogsette, pers. comm.), and best meet the above listed requirements of an ideal insect marker. Fluorescen t dusts can be applied directly to adult flies, or the flies can self mark (auto mark) themselves (Hogsette 1984) Auto marking is accomplished by dusting a known breeding site or by dusting flies in the pupal stage. Adult flies automark themselves by direct contact with the breeding substrate. Emerging flies will auto mark themselves during emergence as the ptilinum typically is dusted while emerging. However, detection of fluorescent dust on the ptilinum is more difficult and time consuming than detection of fluorescent dust on the exoskeleton of flies that were marked as adults, because the ptilinum folds into the head capsule after emergence. Therefore, detection of fluorescent dust on the ptilinum involves squeezing the

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37 head capsule. Additionally, the quantity of dust on the ptilinum is typically much less than that found on flies that are dusted as adults, furthe r increasing the difficulty of using automarked pupae. Fluorescent dusts come in multiple colors which permits selection of colors that are best for working with a particular insect. Day Glo arc yellow and corona magenta are the easiest to use and detect with house flies (Hogsette, pers. comm.). Fluorescent dusts are sometimes visible to the unaided eye, particularly if a great quant ity is present on the insect, but these dusts are best observed under longwave ultraviolet (UV) light. Flight and Dispersal Flight is a dominating characteristic of most adult insects (Johnson 1969). Therefore, any attempt to understand insect influence on disease transmission should incorporate learning about the flight behavior and dispersal and/or migratory tendencies of the target insect. Insect dispersal and migration is reviewed thoroughly by Johnson (1969). Insight into insect populations is provi ded by Southwood (1966); his work has been revised so that newer editions are also available. Methods to quantify insect movement are presented by Turchin (1998). Pedigo and Buntin (1994) compiled the work of several authors to present a detailed resource for various sampling methods applicable to agricultural, including livestock, insects. Flight and dispersal behavior specific to house flies and related species of synanthropic flies with similar breeding habits, such as stable flies and blow flies, has b een reported by a number of researchers and varies drastically in individual studies. In rural areas, house flies can disperse 12 km (Broce 1993a ) and have been documented dispersing up to 21 km (13 mi) (Bishopp and Laake 1921, Alam and Zurek 2004) from their breeding sites. In urban communities, most flies disperse within 1.7 km (1 mi) of release sites (West 1 951, Quarterman et al. 1954, Schoof and Siverly 1954bb; Hanec 1956; Sacc 1964, Milio et al. 1988) However,

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38 house flies have been documented to disperse distances up to 33 km in urban environments (Murvosh and Thaggard 1966). House fly dispersal speed has been documented at a rate of 1 km/h for the first 3 4 h, when dispersal occurred as direct flight over a large swampy area and across rivers with 300 500 m widths in 2450 h (Shura Bura et al. 1962). Dispersal distances and recapture rates might be influenced by the type of flies used, i.e., field collected or laboratory reared. Previous studies have indicated that use of field collected flies is more representative of dispersal under natural conditions than flies that are reared for multiple generations in the laboratory. Eddy et al. (1962) recaptured a 10fold higher percentage of fieldcollected flies than laboratory reared flies; this implies that laboratory colonies may lose the ability to disperse. Dispersal of house flies increases the potential for transmission of zoonotic pathogens to urbanresiding humans, particularly from sites conducive to fly breeding such as dairies (Kaufman et al. 2005, Ahmad et al. 2007, Conn et al. 2007) beef cattle feedlots (Skoda et al. 1993, Thomas 1993, Baldwin et al. 1996, Sanderson et al. 2006), swine facilities (Rosef and Kapperud 1983, Halverson 2000) and poultry facilities (Hald et al. 2004, Watson et al. 2007) If house flies can maintain a travel speed of 1 km/h for an extended period of time, and if house fly dispersal fli ght occurs in a straight line from a breeding site, then house flies could potentially transmit infectious pathogens as far as 12 km in only 12 h (Meerburg et al. 2007). The house fly readily utilizes decomposing fecal/organic matter as well as human food, and moves freely between the two. In locations where dairies and human communities are in close proximity, house fly dispersal between the two could facilitate the transmission of enteric bacteria to humans. External contamination of house flies can range from 2.5 to 29.5 million bacteria per fly (Hawley et al. 1951), and some bacteria can survive up to 3.5 d on the surface of house flies (Peppler 1944). Bacterial contamination of house flies can also occur after flies contact food

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39 crops that have been fer tilized with liquid slurry or solid fecal waste (Islam et al. 2005). Mechanical transmission of bacteria by house flies has been well established by many researchers (Echeverria et al. 1983, Fotedar et al. 1992, Sasaki et al. 2000, Alam and Zurek 2004, Bum a et al. 2004, Ahmad et al. 2007, Nmorsi et al. 2007), whereas, biological transmission also appears likely if E. coli O157:H7 is capable of replicating within the house fly gut (Hawley et al. 1951, Petridis et al. 2006). Fly dispersal has been measured by many types of mark, release and recapture studies using fluorescent dusts, sticky traps, and UV lights (Hogsette 1983, Osek 2001). One of the easiest and most efficient techniques for marking and releasing of large numbers of small insects is the application of fluorescent dust (Hagler and Jackson 2001). Insects such as house flies are collected in the field or mass reared in the laboratory, marked for future identification, released in the field, and recaptured at various distances from their release site. Mark and Release Techniques Many types of markers have been used effectively in the past, but are no longer recommended or are sometimes prohibited under existing regulatory legislature, due to human, animal and/or environmental health concerns. For example, radioactive phosphorus (32P) has been added to adult fly laboratory diets; after successful feeding, 32P labeled flies were released and recaptured in the field (Lindquist et al. 1951, Yates and Lindquist 1952, Eddy 1962, Shura Bura et al. 1962). Marked flies were subsequently counted using readings on Geiger counters (Hoffman and Lindquist 1951, Lindquist et al. 1951, Yates and Lindquist 1952, Eddy 1962, Shura Bura et al. 1962). Hoffman and Lindquist (1951) reared flies in media containing 32P to compare th e efficacy of this method against application by ingestion, and determined that feeding 32P to adult flies was both more effective and cost effective. Lindquist et al. (1951) compared marking adult house flies by adding 32P to the diet against dusting with fluorescent

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40 dusts. They determined that marking with dietary 32P was more efficient and less labor intensive, and they observed that the dusts wore off within 48 h so that identification of dusted flies was difficult. Similarly, Eddy et al. (1962) conclud ed that ingestion of 32P was a more useful marking method than fluorescent dusts, because wild flies had natural fluorescence that was easily confused with the fluorescent marker used in their study. Although there is no universal method of marking insect s, dusts are possibly the most frequently used external markers, due to their ease of use in both application and observation, as well as their low cost, ready availability, and low toxicity. Fluorescent dusts, in particular, DayGlo powdered pigment dust s, have been used to track dispersal and population dynamics, without any observed adverse changes to insect behavior or mortality (Hogsette 1983, Hogsette 1984, Kristiansen and Skovmand 1985). Fluorescent dusts also offer potentially long term investigati ve study possibilities, because the dust has been shown to last up to 3.5 mo in the field (Taft and Agee 1962). Flies are dusted with fluorescent powder, released, and recaptured; subsequent UV light examination of recaptured flies illuminates any retained fluorescent dust on areas of the body that the fly has difficulty grooming. Flies dusted as adults will typically retain dust particles on portions of the thorax; when the puparia are dusted, flies emerge, crawl through the dust, and retain it on their pt inilum (Hogsette 1983). An additional advantage of fluorescent dusts is that their visibility is greatly enhanced when examined under longultraviolet (UV) light. Thus, large numbers of recaptured house flies on sticky traps can be examined rapidly and ea sily under UV light to determine how many are marked. This eliminates time and labor intensive observation methods used with alternative marking techniques, as there is no need to destroy individual insects to observe internally expressed dyes, to apply solvents, or to perform genetic analysis. Application of fluorescent

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41 dusts is relatively easy, inexpensive, and less labor intensive than other insect marking techniques, and enables marking of thousands of insects simultaneously (Zhao et al. 1999). Additio nally, application of fluorescent dusts can be accomplished using mechanical dusters (Hogsette et al. 199 3). Population Dynamics: Monitoring House Fly Populations Fly populations at dairy barns typically need to be monitored in order to determine the effe ctiveness of control methods or to measure seasonal and weather related fly population changes. There are many different methods available to monitor fly populations; each has its strengths and limitations. Ideally, a fly monitoring method will provide sen sitive, accurate results, be easy to use and interpret even by inexperienced persons, be inexpensive, require very little investment of time and labor (Pickens et al. 1972) be protected from cat tle (Morgan and Pickens 1978) and not interfere with cattle operations or endanger cattle health and safety. Monitoring devices must be placed out of reach of cattle, particularly if they are small enough to be eaten by cattle. This is particularly true o f spot cards, described further below. Fly monitoring can be conducted either by passive or active methods. Passive monitoring methods Passive methods of monitoring dairy barn fly populations are those which use one or more of the following: spot cards, s ticky traps, ultraviolet (UV) light traps, and baited traps (Morgan and Pickens 1978) Spot cards are plain white cards (8 x 13 cm) that are conveniently sized for easy transport and use (Axtell 1970, Lysyk and Axtell 1986), but they alternatively may be of larger dimensions such as 13 x 20 cm cards (Pickens et al. 1972). Standard office supply index cards are often used as spot cards. Regardless of size, spot cards are placed vertically or horizontally flush against barn walls, beams, or rafters (Lysyk and Axtell 1985), in areas where house flies rest. Spot cards offer many advantages for monitoring of fly populations. They are

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42 convenient, economical, easy to transport, readily available, and can be used by relatively untrained personnel, including busy far mers. Flies continuously regurgitate and excrete while resting, so spot cards provide a convenient method to obtain house fly relative abundance data in dairy barns when placed in house fly resting areas. Spot cards provide useful information about the changes in house fly populations from week to week, and help ensure optimal fly control efforts, because the success of fly treatments can readily be ascertained. If the number of house fly spots decreases, then the treatment decreased the fly population. Successful treatment would be defined by reducing the number of spots per card below some pre defined threshold that often is farm specific. Spot cards may also be suspended or hung vertically from barn rafters, however, this does not appear to provide usef ul results as Pickens et al. (1972) did not find spot cards sensitive to changes in house fly population density. They observed that the number of fecal and regurgitation spots remained relatively the same despite an artificially doubled house fly populati on due to releases of marked flies within an enclosed barn. However, they hung their spot cards vertically from the ceiling rather than placing them flush against the wall as reported by Lysyk and Axtell (1985). House flies prefer to rest along straight edges. Therefore, hanging large cards from the ceiling was probably not ideal, especially since Pickens et al. (1972) described the presence of a large exhaust fan at one end of the barn. Presumably, the fans created air currents that might have inhibited fly resting on their spot cards, particularly if their spot cards were moving in an air current. Furthermore, Pickens et al. (1972) released laboratory reared flies, and there is some evidence that laboratory reared flies do not take flight as readily as fli es from natural populations (Eddy et al. 1962). Thus, the flies used in their experiment might not have dispersed throughout the enclosed dairy barn as readily as native flies.

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43 In addition to monitoring house fly population changes when placed correctly, spot cards also might permit isolation of fecal and/or regurgitation spots for microbial or genetic analysis of digestive tract contents, although they do not appear to have been used for this purpose yet. In dairy barns that have multiple species of flie s, fecal and regurgitation spots could be due to any of the various species present. However, judicious placement can reduce species overlap by placing cards where house flies are observed resting. Because house flies are not actually captured on spot card s, individual fly species cannot be identified from spot cards using standard methods. However, if fecal and vomit spots contain any host cells from the flies, then the potential exists for identification of particular fly species by genetic analysis such as polymerase chain reaction (PCR). Feces are typically surrounded by a layer of peritrophic membrane and excreted as discrete fecal pellets. Therefore, host cells should be present with the fecal spots, so that the potential for identification of fly sp ecies by analysis of spot cards offers an interesting and promising possibility. Sticky cards (Hogsette et al. 1993) provide another useful house fly population monitoring method. Like spot cards, they are easy to transport, easy to use by untrained perso nnel, and inexpensive. Fresh cards are recommended to achieve optimal results, because the adhesive coating can dry out over time. An advantage that sticky cards offer is the ability to easily identify captured flies to species. Flies become stuck on the adhesive and are therefore readily available for examination, either in the field or in the laboratory. Like spot cards, sticky cards offer the potential for genetic analysis. Because the entire fly body is present, genetic analysis could be used to determi ne the genetic makeup of the flies (for phylogenetic studies) or to analyze the gut contents. Although sticky cards do not appear to have been used for either purpose, they potentially provide another tool for increased understanding of house fly

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44 population dynamics as well as determination of microorganism survival on and within the house fly. Preuss (1951) obtained viable Salmonella sp. from a house fly seven days after it was killed, so the potential for isolating bacteria from house flies captured on st icky cards is strong. A disadvantage of sticky cards is that they can become ineffective if conditions are dusty, because the adhesive becomes covered with dust and debris, so that flies do not adhere after landing on the card. Attempts to correlate spot c ard and sticky counts have been variable. Turner and Ruszler found no correlation, while Geden et al. (1999) found a high correlation. Originally used to capture and monitor stable flies, alsynite traps (Williams 1973) have proven effective for capturing and monitoring house flies and other species of insects. The fiberglass panels reflect sunlight in plane polarized ultraviolet wave lengths, making them attractive to many flying insects: e.g., stable flies (Williams 1973), alate red imported fire ants, Solenopsis invicta Buren, (Milio et al. 1988), and house flies (Geden 2006). The original alsynite trap consisted of translucent rectangular fiberglass panels coated with an adhesive (Williams 1973). Berry et al. (1981) modified the trap by placing adhesive coated sleeves on the interlocking panels instead of applying adhesive directly to the alsynite panels. The sleeves can be removed and taken to a laboratory where insects can be examined and counted. Alsynite traps have proven to be effective, easy to tran sport, and easy to use in the field. A modified alsynite trap (Broce 1988) consists of a translucent rectangular fiberglass panel wrapped to form a cylinder with an adhesive coated clear plastic sleeve that wraps around the outside of the cylinder for easy removal and replacement in the field. Active monitoring methods Active methods of monitoring house fly populations include visual counts, Scudder grids (Scudder 1947, 1949; Pickens et al. 1972), and sweep netting (Morgan and Pickens 1978) Nonbiting fly population den sities can be estimated for some species by active visual counts. For

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45 example, Fannia canicularis (L.), the lesser house fly, has a behavioral habit of hovering in sunny areas. Therefore, one can estimate a lesser house fly population by estimating the num ber of flies hovering in a sunlit circular field 2 m in diameter, by counting in increments of 10 (Morgan and Pickens 1978). Visual observations invariably are estimations, and can be biased due to researcher experience and ability to recognize house flies versus similarappearing muscids. A disadvantage of visual observation is that flies cannot be identified, sexed or aged, so that the population dynamics remain unknown. Scudder grids (Scudder 1947, 1949) are another active monitoring method. They provi de instantaneous snapshot pictures of house fly population densities, and are very useful for monitoring changes in activity levels and population densities. Scudder grids are particularly effective for monitoring the success of control efforts if used r epeatedly at the same physical location both before and after treatment (Scudder 1947). Placement of the grids is critical. Because house flies are not distributed randomly throughout a dairy, but aggregate at food and breeding sites, Scudder grids should be placed in areas of highest fly density. Scudder (1947) found that results were most consistent with observed population trends and most consistent from week to week when he obtained multiple counts at each site and recorded the average of the highest three counts. Some disadvantages of the Scudder grid are that researcher proximity, especially if casting a shadow over the grid ; or repeated placement of the grid in one spot within a short period of time, which can decrease Scudder grid counts, because th e flies move away from the disturbance. Due to fly responses to this monitoring method, Scudder grid counts do not necessarily correlate to an observable population of house flies for the purposes of quantifying population densities. Grids should be placed by the same person to decrease user induced variability influences on fly counts. As with visually observed flies, flies that land on Scudder

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46 grids cannot be identified, aged or sexed, so information regarding the population dynamics remains unknown. Howe ver, for the experienced person, Scudder grids can provide an inexpensive, easy to use and reliably repeatable method for monitoring increases or decreases in fly populations. Their ease of use and portability in the field make them useful monitoring metho ds at dairies and other animal rearing facilities. Sweep netting at dairies is another fast and easy method used to monitor adult house fly populations. Like Scudder grids, sweep netting is performed onsite to gain an instantaneous snapshot picture of house fly and other flying insect population densities (Dhillon and Challet 1985, New 1998). However, unlike the Scudder grids, repeated sweep netting reduces the available population, because one is removing individuals from the environment and greatly di srupts flies in a given area. In contrast, Scudder grids do not directly impact population sizes and minimally impact flies in a given area. Sweep netting would not noticeably impact highdensity populations; however, when fly populations are low, sweep ne t counts for successive sweeps may decrease with each sweep. Like the Scudder grid counts, sweep nets should be performed in fly aggregation areas, at the same time of day, and by the same person. Sweep net counts can vary dramatically between collecting i ndividuals, based on experience and technique. It might be important to note that sweep netting captures only airborne flies, and flies are typically captured by purposely disturbing adult flies that are resting, feeding or ovipositing. Therefore, sweep ne tting might not provide accurate information regarding the normal level of airborne activity. However, an advantage of sweep netting over other monitoring methods is the capture of live adults, so that analysis of population ratios can be made. Collected f lies can be identified to species, sexed and aged and female flies can be examined for reproductive status. Collection of mostly young adults 1 3 d old would provide information that the flies had only

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47 recently emerged. Because sweep netting enables the researcher to obtain information regarding population dynamics, it is a useful monitoring method at dairies and other animal rearing facilities. House Fly Economical Impacts and Disease Outbreaks House flies are strongly associated with humans and livestoc k. This synanthropy, when combined with the house fly's caprophagic and saprophagic eating habits, makes the house fly an important vector of human diarrhegenic pathogens. Diseases which are transmitted via the fecal oral route are especially prone to diss emination by the house fly. In locations where food is being prepared for consumption, such as restaurants, the potential for bacterial contamination by the fly is tremendous, as the fly travels freely between decomposing organic matter found in restaurant garbage dumpsters, exposed kitchen surfaces and foods, dining tables, and even restaurant bathrooms. The bacterial diversity and quantity on restaurant associated house flies has been examined (Nayduch et al. 2001, Butler et al. 2010), especially at readyto eat (RTE) food establishments (Macovei and Zurek 2008). Diarrheal diseases impart an enormous economic toll on human and agricultural animal (e.g., cattle) populations, with severe health and economic impacts. Hospital expenses for human patients with infectious diarrhea can four times greater than for other patients. Similarly, medication expenses can be four times higher and the length of hospitalization can be three times longer (Suda et al. 2003). Economic impacts include lost income for families t hat must miss work, as well as lost profits for employers (CDC 2002). For cattle, increased operating expenses are incurred by dairy farms, feedlots and cattle rendering plants that must increase fly and pathogen surveillance and management measures to com ply with federal regulatory mandates (CDC 2009). Indirect losses include increased labor costs associated with fly control and sanitation efforts, which may include increased fuel costs to operate composting and waste-

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48 removal equipment, increased water costs to wash barns and clean equipment, increased expens es for insecticides and traps, and additional wages to employees who perform all the additional tasks (Lazarus et al. 1989). It is interesting to note here that expenses re lated to insecticides can incr ease quickly because of the house flys rapid development of insecticide resistance. House fly insecticide resistance can occur quickly enough to necessitate increased usage of insecticides, thus driving up the expense dramatically. Alternatively, one may switch to a pesticide containing a different activeingredient; however, this still results in overall increased expenditure towards the effort of controlling fly populations in an effort to prevent diarrheal disease occurrence. Additional indirect losses can include legal expenses and forced farm closures (Thomas and Skoda 1993) Courts can impose stiff fines upon many different foodrelated industries including producers, distributors, and restaurants. A recent example: in June, 2008, a lawsuit led to a $13.5 million settlement after a child in Milwaukee, Wisconsin died due to consumption of E. coli O157:H7contaminated food (Powell, 2008). House Fly Management Pest control efforts include cultural, mechanical, biological, and chemical methods. Each metho d is discussed below, and the integration of these methods into one comprehensive control program known as Int egrated Pest Management (IPM) Cultural c ontrol Cultural control includes sanitation and management efforts, e.g., moisture and manure removal at dairies designed to prevent fly populations from building to unacceptable levels (Stafford 2008) by eliminating immature fly development areas Flies breed in spilled feed moist hay and manure Weekly removal of these materials is recommended (Rutz et al 1994). S pilled feed and manure can accum ulate and retain moisture under fence edges, along the sides of liquid

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49 manure pits, and in other hardto reach areas ( Rutz et al. 1994, Farkas and Hogsette 2000). Preventing moisture in fly breeding areas, e.g., in spilled grains, is an important part of cultural control (Farkas and Hogsette 2000). Watering devices should be maintained to prevent leaks, and manure should be removed to maximize drainage. Flies can breed in manure that lines manure pits or floats on the surface of manure pits and lagoons. Therefore, cultural control of fly breeding sites includes preventing manure clumps from lining manure pits or from floating on the surface (Farkas and Hogsette 2000) W ithout adequate removal of manure and other fly breeding sources, chemical e fforts to control th e house fly population will be less effective and more expensive (Farkas and Hogsette 2000). C ultural control methods also encourage the growth and development of house fly predators and parasites (biological control). Additional cultural control methods include using screens as exclusion barriers on windows and doors, preventing access to garbage, and composting garbage manure and soiled bedding properly so that decomposition is aerobic and hot enough to ki ll developing flies (Stafford 2008). Mechanical c ontrol Mechanical control methods use non chemical devices to kill flies that are in the environment. Fly swatters provide low tech mechanical control of individual house flies (Stafford 2008). Low numbers of flies can be controlled by using sticky traps, jug or cylinder traps and bag traps. Sticky traps are coated with adhesive materials, so they are less effective when fly numbers are high or when environmental conditions are dusty (Stafford 2008). Sticky traps are most effective when used indoors; however, they cannot be used in foodpreparation settings because become unsightly and they can drip when temperatures become too warm (Carlson and Hogsette 2007). Fluorescent or ultraviolet light emitting electrocution traps are useful for mechanically controlling larger fly populations (Stafford 2008). The flies are

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50 electrocuted when they contact the electric mesh. However, electrocution of insects results in airborne scattering of insect parts and infectious m icroorganisms (Pickens 1989, Ananth et al. 1992, Broce 1993b, Tesch and Goodman 1995, Broce and Urban 1998). Therefore, use of electrocuting traps is not recommended, and is sometimes prohibited, in medical and foodpreparation settings. An additional disa dvantage of electrocuting grids is that they kill high numbers of nontarget insects (Frick and Tallamy 1996). Many mechanical traps use ultraviolet light traps with sticky glue boards instead of electrocuting grids, although the attractiveness of these tr aps relative to the attraction of food in the foodpreparation area is unknown (Carlson and Hogsette 2007). Bulbs in light traps lose their effectiveness over time, so that they become less attractive to flies. Thus, it is important to replace bulbs on a t imely basis. Additionally, indoor traps should be installed so that they attract flies that are already in the local environment, rather than attracting flies from far away into the establishment. Some mechanical control traps use attractant baits that d raw house flies into a nontoxic solution (Stafford 2008). Examples include jug traps, bag traps, and metal or plastic cylinder traps. Baited traps are useful for controlling large numbers of house flies; some use only water as the bait, while others add proprietary chemical mixtures that mimic natural food materials or pheromones, such as the female house fly sex attractant (Z) 9tricosene. Because (Z) 9 tricosene is odorless to humans, it is a useful indoor bait. In contrast, baits that mimic natural food sources typically have strong unpleasant odors and are therefore best suited for outdoor use. Bait traps come in a wide variety of styles and sizes, with some designed to be hung and others designed to be placed on the ground. In general, bait traps take advantage of the house flys positive phototrophic behavior by providing small access holes through which the flies can enter the trap, but not exit. Therefore, the flies die inside the trap. Jug and cylinder (container) bait traps can be

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51 reused by emptying them when approximately one third full, and rebaiting, so that they provide an economical control method. Homemade bait traps are easily made from milk cartons and plastic soda bottles by cutting the top portions off and replacing them in an inverted pos ition to acts as a funnel. Flies will readily enter the carton or bottle through the funnel, but will not exit. Like bag traps, carton and bottle traps are disposable (Stafford 2008). Biological c ontrol Biological control of house flies is achieved through predation or parasitism by natural house fly enemies or by infection with pathogens that kill the house flies While biological control occurs unassisted in the natural environment, attempts to amplify its actions through release of increased bio logical control agents are ongoing. Natural enemies of house flies include other arthropods, fungi, bacteria, and viruses (Barnard 2003, Geden 2006, Stafford 2008). Parasitoid wasps used for biological control of house flies are often species in the family Ptero malidae, from the following genera: Spalangia, Muscidifurax and Nasonia ( Morgan et al. 1979, Crespo et al. 1998, Tobin and Pitts 1999, Floate et al. 2000, Kaufman et al. 2001a, Geden 2006, McKay et al. 2007 Birkemoe et al. 2008 ). One benefit of the paras itoid wasps is their relative targetspecificity, due to the postulate d co evolution of the parasitoids with their synanthropic muscoid hosts (Pimentel et al. 1963). Some species of wasps will target both house flies and stable flies, which are two of the most economically damaging fly species in livestock environments. Efforts to identify new species of parasitoid wasps and to commercially rear them are ongoing. Six hymenopteran parasitoids that specifically attack fly pupae were evaluated by Geden (2006): Muscidifurax raptor Girualt and Sanders, Spalangia cameroni Perkins, Spalangia nigroaenea Curtis, Spalangia endius (Walker), Spalangia gemina Boucek (Hymenoptera: Pteromalidae), and Dirhinus himalayanus (Hymenoptera: Chalcididae). Hogsette et al. (2001)

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52 obtained seven hymenopteran pupal parasitoids from house fly pupae in Hungary: Spalangia cameroni Perkins, S. nigroaenea Curtis, S. endius Walker, Pachycrepoideus vindemiae Rondani, Trichomalopsis sp., and two undetermined spp. of Diapriidae. Other types of wasps reported as parasitoids of house flies include species in the family Ichneumonidae: for example, Exeristes comstockii (Cress) (Hymenoptera: Ichneumonidae) (Bracken 1965). Hogsette et al. (2001) reported an undescribed species of Brachymeria parasi tizing a house fly pupa in Hungary. Some dipteran species are also useful for biological control of house flies. Some are predators, such as the bronze dump fly (Byrd and Castner 2009), sometimes called the black garbage fly, Hydrotaea (=Ophyra) leucostom a (Weidemann) (Diptera: Muscidae) (Anderson and Poorbaugh 1964), and the black dump fly, Hydrotaea (= Ophyra ) aenescens ( Weidemann) (Diptera: Muscidae) (Hogsette and Jacobs 1999 Hogsette et al. 2002). Hydrotaea larvae actively pursue and attack house fly larvae, but rarely the pupae (Anderson and Poorbaugh 1964) Hydrotaea eat the visceral tissues, but not the larval integument or puparium One Hydrotaea fly can kill up to 20 house fly larvae in one day (Anderson and Poorbaugh 1964) Hogsette and Washington (1995) developed a method to mass rear Hydrotaea aenescens for biological control studies. Other Diptera are effective biological control agents by nonpredatory methods. One such example is the black soldier fly, Hermetia il lucens (L.) (Diptera: Stratiomyii dae) (Furman et al. 1959; Sheppard 1983; Sheppard et al. 1994; Sheppard et al. 2002). Studies indicate that soldier fly larvae outcompete house fly larvae for food resources in the breeding habitat, and, in this manner, lim it house fly growth and larval development (Sheppard 1983) Bacteria such as Bacillus thuringiensis (Zho ng et al. 2000, Ruiu et al. 2007) and Brevibacillus laterosporus (Ru iu et al. 2006) have been reported as useful for biological control

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53 of house flies caterpillars and mosquito larvae (Swadener 1994) due to production of insecticidal endotoxins ( Zhong et al. 2000; Ruiu et al. 2006, 2007) B acillus thuringiensis subsp. israelis ( Bti ) produces a delta endotoxin ( Zhong et al. 2000, Ruiu et al. 2006). However, t here is evidence that Bti and B. laterosporus are toxic against the house fly parasitoid, M. raptor although much less so than against the house fly (Ruiu et al. 2007) Fungi are considered by some researchers to show promise as biological contro l agents against house flies (Mullens and Rodriguez 1986, Geden et al. 1993, Steinkraus et al. 1993, Watson and Petersen 1993). Two fungi that successfully infect and kill adult house flies are Entomophthora muscae (Chon) Fresenius and Beauveria bassiana ( Balsamo) Vuillemin However, these fungi are limited to specific climatic conditions, and are dependent upon high fly population densities (Watson and Petersen 1993) Although t hese fungi difficult to produce commercially well enough to be effective and ec onomically viable commercial house fly biological agents ( Geden et al. 1993, Watson and Petersen 1993) recent rearing techniques have resulted in a commercial product that is highly effective (Kaufman et al. 2005a) Beauveria bassiana has proven to be effective against both adult and larval house flies (Watson et al. 1995). Chemical c ontrol In general, chemical control of house flies has been the most widely used approach over the past 60 years. Ins e c ticides are applied in a variety of ways, including spa ce sprays, residual wall sprays, feed through products, onanimal applications, misting systems and toxic fly baits (Rutz et al. 1994) Pyrethrin fogs and space sprays are recom m ended for initial use because they work well in conjunction with biological control for integrated pest management (Kaufman 2002). Insecticides can adversely affect bi ological control agents, particularly if used early in the season because parasite populations lag behind house fly populations (Rutz et al. 1994). Elimination of biol ogical control agents can result in an increased dependence on insecti ci des

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54 that can increase the development of insecticide resist a nce in the house fly population (Rutz et al. 1994). R esidual pesticides are recommended only for use emergencies and for use late in the season (Kaufman 2002). Toxic fly baits cannot be used in food establishments, including milk storage rooms on dairies, or in areas where children are present. They also should be avoided in areas frequented by children or pets, and must be pla ced in a container but not sprinkled on the ground. Resistance to chlorinated hydrocarbons develops more quickly in larvae than in adults (Miles 1959). He concludes that chemical control should therefore be restricted to adults, to minimize development of resistant fly strains. However, adult flies can develop tremendous resistance against over used pesticides: Cao et al. (2006) reported 13to 250fold greater deltamethrin resistance in adult house flies collected from urban garbage dumps in Northern Chin a than from laboratory susceptible strains. Integrated Pest Management Integrated pest management i ncorporates a combination of several available fly control methods (Kaufman et al. 2002) Each control method that is used to control fly populations at ani mal rearing facilities such as dairies is insufficient or uneconomical if used alone. The foundation of any successful IPM program is sanitation and manure management. Without adequate cultural controls, neither biological nor chemical controls will be eff ective. Use of b iological control decreases dependence upon chemical insecticides (Farkas and Hogsette 2000). However, natural parasitoids used for fly control differ in efficacy depending upon geographical region, season, climate, habitat host density and host distribution. Thus, it is important to use naturallyoccu r ring species of parasitoids for a particular area, and it is vital to understand both the house fly and parasitoid bionomics to effect a successful longterm biological component of the overa ll IPM program (Farkas and Hogsette 2000) P arasitoid

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55 populations can be reduced below effective levels by indiscriminate use of broad spectrum insecticides used to control house fly populations, and by manure removal. Mass releases of parasitoid wasps are often necessary to reduce house fly populations throughout the season, because the parasitoids need to be consistently present in the manure. For this reason, although sanitation is the first step in successful fly control, manure removal should not be complete, but should leave a base of old manure for house fly parasites and predators (Farkas and Hogsette 2000) Chemical control can achieve a rapid reduction in adult flies, but is limited in use due to side effects or poisoning when overused, to breed se nsitivity, to interactions with medicatioin administered to livestock animals, to contamination of food products such as milk, and to the development of resistance among the fly populations (Farkas and Hogsette 2000, Pimentel 2002). Therefore, chemical con trol should be used in conjunction with cultural/sanitation, mechanical, and biological control methods (Rutz et al. 1994, Farkas and Hogsette 2000, Kaufman et al. 2002). Usefulness of H ouse F lies House flies are useful in some ways that might be unexpected. For example, house fly pupae have been used to achieve relatively speciesspecific chemical control of several ant species, including Solenopsis invicta Buren, the red imported fire ant (RIFA). Ants are extremely effective predators of house fly larva e. Williams et al. (1990) administered hydramethylnonand fenoxycarbcoated house fly pupae to eight species of ants. They achieved 100% mortality in three species (including RIFA) and <20 % mortality in the remaining five species by using house fly pupae as bait carriers. They note that use of house fly pupae permits more selective control over species that consume house fly pupae, and that mass production of the house fly pupae could make this potential method of ant control more economical (Williams et al. 1990). For

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56 baits, house fly pupae was not practical. However, house fly pupae treated with cyclodienes were used for aerial applications in parts of the United States. House flies are potentially useful in an interesting agricultural application to he lp prevent browning of apples. House fly pupae contain high levels of an apple polyphenol oxidase inhibitor that inhibits adverse browning in apple and apple products (Yoruk et al. 2003). The house fly might be a promising livestock and poultry food resource (Eby and Denby 1978, Boushy 1991) due to the very aspects of the house flys biology and behavior that make it a global pest. The house fly converts decomposing organic matter and fecal materials into body tissue during its larval stages. In doing so, it reduces waste, generates energy and creates a high nitrogen food resource. House flies have been used as feed supplements for swine (Poluektova et al. 1980, Chiou and Chen 1982) and poultry (Calvert et al. 1969, Papp 1974, Teotia and Miller 1974, Gawaad and Brune 1979). Freeze dried fly larvae are also commercially available for use as fish feed. However, the use of immature flies as feed supplements might not be advisable unless they can be subjected to bactericidal conditions, because there is evidenc e that animals can become infected with pathogenic bacteria present in the consumed flies (Gerberich 1952). Even if flies are not used as a feed resource, they are still useful because they perform the beneficial act of decomposing manure (Miller et al. 1974, Beard and Sands 1973). This could reduce the amount of waste and decrease labor costs associated with wastemanagement efforts. Although not intentionally provided as supplemental feed, adult flies might be consumed by animals, particularly poultry, a fter they are killed by pesticide treatment at an animalrearing facility. Therefore, efforts to reduce consumption of flies killed by pesticides might be advisable. Greenberg (1959e) postulates that consumption of adult flies might result in pathogen

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57 tran smission after citing the work of Gross and Preuss (1951) in which they determined that viable Salmonella sp. were recovered in flies 7 d after they were killed with DDT. Survival of pathogenic bacteria in dead insects illustrates the importance of sanitat ion in conjunction with fly control in order to reduce disease transmission potential at animal rearing facilities. Enterobacteriaceae Enterobacteriaceae Rahn 1937 is a family of opportunistic, facultative anaerobic, non spore forming, gram negative, rodshaped bacteria, found as normal commensal fauna in healthy human and domestic animal gastrointestinal tracts (Caprioli et al. 2005, Madigan and Martinko 2006, FDA CFSAN 2007a) Enteric microbiological communities tend to be very complex, with bacterial populations of more than 1014 cells, which is ten times more cells than the number of cells that constitute the human body (FDACFSAN 2007a) Enteric bacteria belong to more than 500 different species (Steinhoff 2005) which collectively perform various metabolic activities, such as digestive fermentation and vitamin synthesis upon host consumed nutrients (FDACFSAN 2007a) With this level of complexity, resultant fecal bacteria concentrations can exceed 1012 cells/g of feces (Couteau et al. 2001) Some enterobacteria are opportunistic pathogens that cause diarrhea, dysentery, meningitis, typhoid fever, and food poisoning (MEDIC 1995) to mammalian hosts through a variety of mechanisms (Manafi 2003) Disease causing enterobacteria include several closely related genera: Escherichia, Shigella, Salmonella, Yersinia, Klebsiella, Pr oteus, Edwardsiella, Citrobacter, Enterobacter, Serratia, Providencia, and Morganella (FDACFSAN 2007a) Although Escherichia is listed at the genus level with the above group, i t is unique due to the fact that only one of its seven species (Euzeby 2008, NCBI. 2008) E. coli is known to be pathogenic to mammals. In contrast, other genera have several pathogenic species.

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58 In addition to shared morphological and physiological traits, members of the Enterobacteriaceae share biochemical characteristics t hat make differentiation and identification difficult: all ferment glucose, reduce nitrates, and are oxidasenegative. Despite these similarities, other biochemical characteristics are diagnostic for differentiation. For example, lactose fermentation is ty pical of nonpathogenic enterobacteria, while nonfermentation of lactose is typical of pathogenic enteric bacteria. Selective and/or differential media used in the laboratory to identify pathogenic strains takes advantage of this lactose fermentation char acteristic by inclusion of lactose and dyes which provide presumptive identification by easily observed color changes where fermenting and nonfermenting bacteria will produce colonies of distinctly different colors. Among the Enterobacteriaceae, the genus Escherichia is comprised of seven species (Euzeby 2008, NCBI 2008) one of which is E. coli Pathogenic strains of E. coli present health risks, partially because of their low infectious dose, which is variously reported as less than 100 organisms/g of f eces (USDA FSIS 2008a, USDA FSIS 2008b) less than 50 organisms/ g of feces (Tilden et al. 1996), and possibly as few as 10 organisms/g of feces (FDA CFSAN 2007b). Escherichia coli Escherichia coli was originally described in 1885 as Bacterium coli commune, by Theodor Escherich ( Janda and Abbott 2006, NCBI 2008, Todar 2008) An additional synonym of E. coli used in older literature is Bacillus coli Migula 1895 (NCBI. 2008) Esherichia coli is the most abundant, commensal, facultative anaerobic bacteria in many mammals, including humans. (Donnenberg 2002). Homeothermic animals are the natural reservoir of E. coli and E. coli is part of the normal gut fauna in nearly all domestic animals (Bettelheim 199 1), and accounts for approximat ely one percent of fecal biomass (Janda and Abbott 2006) Human stool contains a diverse bacteria fauna of 1011 cells per cubic centimeter wi th 109 E. coli per cubic centimeter

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59 (Neidhardt et al. 1996). Escherichia coli is a rod shaped bacterium with an approximate length of 2.5 m and diameter of 0.8 m (Berg 2004). In contrast, a human hair is 80 l in diameter (Neidhardt et al. 1996). When pl aced in warm nutrient broths, E. coli replicates in 20 min (Berg 2004). Commensal E. coli appears to produce cofactors and to inhibit pathogenic colonization in the digestive tract (Donnenberg 2002). Escherichia coli is comprised of more than 700 strains (Todar 2008) and more than 200 of these strains produce Shiga toxins (Thorpe et al. 2002), of which most of those strains are pathogenic (Madigan and Martinko 2006, FDA CFSAN 2007a) All pathogenic strains are enteric (Madigan and Martinko 2006) but the pathogenic potential of a particular strain depends on the specific collection of virulence genes (Donnenberg 2002). Within the anaerobic gastrointestinal tract of animals, non pathogenic commensal strains of E. coli synthesize vitamins and inhibit growth of pathogenic microorganisms (FDA CFSAN 2007c). Both behaviors benefit the mammalian host with improved nutritional uptake. In humans, E. coli is the predominant bacterium in the large intestines (FDA CFSAN2007a), whereas in cattle, E. coli is a normal component of the rumen, the largest chamber of the four chambered stomach (Callaway et al. 2004 ) Although consistently present in all animal gas trointestinal tracts, E. coli prevalence, density, survival and fecal shedding rates can differ among hosts. For example, cattle shed E. coli O157:H7 intermittently (Zhao et al. 1995, Shere et al. 1998). Differences in host diet have been implicated in inc reased fecal shedding of E. coli O157:H7. For example, E. coli from grain fed cattle are especially resistant to low pH and have greater survival rates in acidic enteric environments than E. coli from forage fed cattle (Callaway et al. 2003)

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60 Outside the host's facultative anaerobic intestinal environment, E. coli can be outcompeted by other species of bacteria. Host excretion of manure rapidly exposes E. coli to multiple systemic shocks. The two most important shocks relate to transition: 1) from an ideal 37 C to ambi ent air temperature, and 2) from an anoxic environment to a highly oxygenated environment, which is relatively toxic to E. coli (Doyle and Beuchat 2007). Separately, either of these two shocks can result in serious, but reversible, injury to E. coli ; howev er, when combined, they can cause irreversible damage and mortality. Because E. coli is a facultative anaerobe, the aerobic environment is not directly lethal to E. coli Instead, the aerobic environment presents an indirect, potentially more harmful, obs tacle to E. coli 's survival outside of the enteric environment: competition with aerobic microorganisms. Competing organisms can inhibit E. coli directly by secreting antibiotics or toxins (Xavier and Russell 2006), or indirectly by consuming available nut rients (Hibbing et al. 2010). Concentrations of E. coli in hosts differ among host animal species. Within a particular animal, concentrations also can differ depending on gastrointestinal location. In cattle, concentrations are high in the mucosal anal region because it is the site of E. coli O157:H7 attachment (Greenquist et al. 2005). In cattle, prevalence and density of E. coli in fecal samples varies widely, and may be influenced by factors including diversity of the bacteria community within the gastrointestinal tract, diet, season, lactating stage (Hancock et al. 1994), and application of vaccines (Van Donkersgoed et al 2005). Many different strains of E. coli may be present in one individual simultaneously, as seen in humans (Beutin et al. 2004) and cattle (Majalija et al. 2008).

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61 There is some level of host specificity between certain strains of E. coli and a particular animal host. While commensal bacteria serve a useful, and essential role in digestion for the host animal, these bacteria can becom e pathogenic if they are introduced into a different animal species. For example, E. coli O157:H7 is a commensal bacterium in both livestock and wild ruminants. However, this strain of E. coli is a human pathogen that can result in severe illness and death. In humans, E. coli O157:H7 causes diarrhea, dysentery, hemolytic colitis (HC), and if left untreated, can lead to development of hemolytic uremic syndrome (HUS) and acute kidney failure (Tarr and Neill 2001). Pathogenic Escherichia coli Although most E. coli are beneficial to their associated host, a subset of E. coli are pathogenic, and can cause diarrheal diseases (FDA CFSAN 2007b). Escherichia coli is a clonal species, with clones differentiated into serotypes based on various combinations of somatic (O) and flagellar (H) antigens (Wang and Reeves 1998). The O and H antigens are most frequently used for serotype identification and differentiation. There are at least 181 E. coli O antigens (Durso et al. 2007). Many of these antigens are associated with pathogenicity (Wang and Reeves 1998) whereby these O antigens are considered virulence factors (Wang and Reeves 1998). There are approximately 200 pathogenic strains of E. coli and they are broadly grouped into large subsets: pathogenic E. coli (FDACFSAN 2007b) enterovirulent E. coli FDACFSAN 2007b), diarrheagenic E. coli (Nataro and Kaper 1998, FDA CFSAN 2007c) and enterohemorraghic E. coli (FDACFSAN 2007b). There i s overlap between some of the groups, because E. coli strains are placed into groups based on possession of particular virulence factors such as O and H antigens and biochemical traits. Therefore, serotypes that possess multiple virulence factors sometimes fall into more than one group. In general, diarrheagenic E. coli are a subset of pathogenic E. coli and enterohemorrhagic E. coli are a subset of diarrheagenic E. coli

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62 Pathogenic E. coli include all of the disease causing strains, regardless of etiolog y. Some of the more infectious and therefore most studied O antigen serogroups include: O26, O55, O86, O103, O111ab, O119, O125ac, O126, O127, O128ab, O142, and O157 (Murinda and Oliver 2006, (FDA CFSAN 2007a, Monday et al. 2007). Some infectious strains of E. coli result in urinary tract infections (FDA CFSAN 2007b), meningitis, or other nondiarrheagenic diseases. However, most pathogenic E. coli cause some type of diarrhea. The primary cause of diarrhea resulting from E. coli infections is due to release of Shiga toxins (CDC 2008), but not all E. coli that cause d iarrhea possess the Shiga toxinproducing genes. Clinical presentation of diarrhea can differ greatly among diarrheagenic strains of E. coli and among patients, so that grouping of strains based on symptoms can be ambiguous or misleading. Therefore, labor atory identification of suspected E. coli bacteria samples should be performed to identify the particular serotype. Enterovirulent strains are those E. coli that can be categorized by virulence factors (VFs), which are unique within each group: enterohemo rrhagic E. coli (EHEC), enterotoxigenic E. coli (ETEC), enteropathogenic E. coli (EPEC), enteroinvasive E. coli (EIEC), enteroaggregative E. coli (EAEC) and diffusely adherent E. coli (DAEC) (FDACFSAN 2007b). Of the enterovirulent E. coli EHEC causes the most foodborne outbreaks, and the primary serotype is E. coli O157:H7. Note that additional serogroups O26, O111, O126, and O103 are nonO157 serogroups which have recently resulted in infectious disease outbreaks ( Food Source 2006). However, the focus of this review is limited to EHEC, E. coli O157:H7, a diarrheagenic and enterohemorrhagic serotype. Diarrheagenic strains include those featuring the clinical symptom, namely diarrhea, that accompanies enteric E. coli infections. However, it should be note d that diarrhea can result from infections of other pathogens, so that diarrhea is not necessarily indicative of E. coli infection. In

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63 fact, diarrhea is not even limited to bacterial infections. For example, most diarrhea in children is caused by two virus es: norovirus and rotavirus (Doyle and Beuchat 2007). Gastrointestinal diseases, regardless of causative organism, generally always result in diarrhea from a sloughing off of the epithelial cells lining the intestine. Enterohemorrhagic E. coli strains ar e a subset of diarrheagenic strains. These disease causing E. coli strains are limited to those that cause bloody diarrhea. Norovirus, rotavirus, and adenoviruses also can all result in bloody diarrhea (Doyle and Beuchat 2007). Within E. coli strains, seve ral serotypes are enterohemorrhagic, particularly those that possess the Shiga toxin producing genes. Some strains of E. coli overlap into other areas of pathogenicity, because they possess multiple virulence factors. For example, although E. coli O55:H7 i s primarily enteropathogenic, it is also enterohemorrhagic. Thus, it is often grouped together with O157:H7 into an "O157:H7 complex" of enterohemorrhagic E. coli strains (Feng and Monday 2005). Escherichia coli O157:H7 Within the E. coli O157:H7 complex, E. coli O157:H7 is an enterohemorrhagic strain that possesses many diseasecausing virulence factors (Janda and Abbott 2006) This pathogen is known as Shiga toxinproducing E. coli (STEC) because it produces Shiga toxins, and is also referred to as enter ohemorrhagic E. coli (EHEC) because it is an enteric bacteria that causes diarrhea and dysentery (Janda and Abbott 2006). Escherichia coli O157:H7 and Cattle Escherichia coli O157:H7 is a zoonotic bacterium that causes human disease. Cattle are the primar y reservoir (Hussein et al. 2003, Davis et al. 2005, Sargeant et al. 2005, Alam and Zurek 2006) although other species of domestic animals reared for food also serve as reservoirs, including sheep (Chapman et al. 1997, Keen et al. 2006) goats (Pao et al. 2005, LeJeune et al. 2006), swine ( Chapman et al. 1997, Keen et al. 2006) and poultry (Chapman et al. 1997).

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64 According to a report issued by USDA:APHIS:VS (2007), recent cattle herd prevalence rates in the United States ranged from 22100%, although preval ence rates for individual animals within the herds ranged from 0 9.5%. Hancock et al. (1997) found the prevalence of O157 lowest in adult cattle (0.4%) and highest in weaned heifers (1.8%). He also determined that E. coli O157:H7 of the cattle gut is trans ient, with fecal shedding lasting a median of 30 d. Infection with E. coli O157:H7 is not limited to livestock and poultry as it has also been isolated from many other mammals and arthropods including deer ( Asakura et al. 1998, Dunn et al. 2004a) opossum (Renter et al. 2004 b) pigeons (Morabito et al. 2001) rabbits (Scaife et al. 2006, Fremaux et al. 2008) house flies (Alam et al. 2004, Sanderson et al. 2006) blow flies (Fotedar et al. 1992) and slugs (Spro ston et al. 2006) Although adult cattle are as ymptomatic carriers of E. coli O157:H7 bacteria (Porter et al. 1997), E. coli O157:H7 can cause disease in unweaned cattle that do not possess a fully developed rumen, making their digestive system comparable to that of humans. In adult cattle, E. coli O15 7:H7 is a commensal microorganism in the rumen, which helps provide dietary nutrients to the host animal (Frandson 1969). Caprioli et al. (2005) reported that poultry and pigs do not serve as EHEC sources; however, other researchers reported contradictory results (Doane et al. 2002). Caprioli et al. (2005) concluded that poultry and pigs become infected by exposure to cattle and other ruminant (sheep, goats, water buffalo, and deer) excrement. Although all ruminants may potentially serve as reservoirs for E coli O157:H7, cattle serve as a dominant reservoir of E. coli O157:H7, and are considered by many as the primary reservoir (Loneragen and Brashears 2005). Feedlot cattle preharvest diets have been modified in various ways in an effort to reduce the leve ls of E. coli O157:H7 in feedlot cattle. Addition of microorganisms to the diet appears to be one of the most promising diet modifications. In

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65 contrast, addition of brown seaweed does not seem to reduce levels of E. coli O157:H7 (Loneragen and Brashears 2005). In addition to modification of diet, administration of vaccines, sodium chlorate, and neomycin sulfate to feedlot cattle has demonstrated beneficial results towards reduced E. coli O157:H7 pre harvest levels. In contrast, addition of chlorine to water sources does not appear to reduce E. coli O157:H7 levels in pre harvest feedlot cattle (Loneragen and Brashears 2005). Escherichia coli O157:H7 Outbreaks The origin of the enterohemorrhagic strain of E. coli has not been identified, although it emerged v ery recently. It was first identified, although not associated with disease, in 1975 by the Centers for Disease Control and Prevention (CDC) (Riley et al. 1983). Between 1978 and 1983, laboratory analysis of stool specimens from six sporadic gastrointestinal cases resulted in isolation of E. coli O157:H7: five of the cases were for patients with clinical symptoms of hemolytic colitis, while the sixth was isolated from a patient with an unknown medical history. In addition to isolation from sporadic cases, E coli O157:H7 was isolated and determined to be the causative agent for three 1982 gastrointestinal outbreaks. The first two occurred in Oregon and Michigan, USA, and caused 47 known illnesses among fast food patrons (Riley et al. 1983) The third outbreak in 1982 resulted in 31 cases, 4 hospitalizations, and one fatality among a population of 353 elderly persons in Ottawa, Ontario, Canada (CDC 1983) Thus, within a very short spa n of seven years, a newly emerged E. coli strain quickly expanded its pathogenic impact upon the human population. Denny et al. (2008) and Feder et al. (2003) cite annual E. coli O157:H7 disease estimates provided by Mead et al. (1999) for the United Stat es. They estimate that approximately 73,000 cases and 61 fatalities occur each year in approximately 500 outbreaks. An estimated 27 40% of stricken individuals progress to either severe hemorrhagic uremic syndrome (HUS) or renal

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66 sequelae. They infer that t he number of illnesses, hospitalizations and deaths due to E. coli O157:H7 may be under reported, due to different reporting mechanisms between states and among doctors. As of 1999, all food borne illnesses, including but not limited to E. coli O157:H7, we re determined to be responsible for as many as 5,000 fatalities, with 325,000 hospitalizations out of a total of 76 million illnesses (Mead et al. 1999, Mai et al. 2006) It is possible that some of the cases of u nknown etiology were due to undetected E. coli O157:H7. Shiga toxin producing E. coli were responsible for an estimated 100,000 cases with 2,000 hospitalizations and 91 deaths (Frenzen et al. 2005 as cited in DuPont 2007). For humans, hospital expenses for patients with infectious diarrhea can be as high as $24,000 versus $9,000 for patients that do not have infectious diarrhea (Suda et al. 2003). Similarly, medication expenses can be four times higher (~$4,000 vs. ~$1,000) and the length of hospitalization can be three times longer (22 vs. 7 days) (Suda et al. 2003) Economic impacts can also include lost income for families that must miss work as well as lost profit for the employers of affected persons (CDC 2002). Shiga toxin producing E. coli were responsible for an estimated 100,000 cases with 2,000 hospitalizations and 91 deaths.(Frenzen et al. 2005 as cited in DuPont 2007). Total numbers of E. coli O157:H7 cases worldwide from 19822006 reflects movement into new geographical regions. During this 24yr period, Doyle et al. (2006) reported worldwide E. coli O157:H7 statistics garnered from reviews of the published literature as: 207 outbre aks and 26,179 cases. The economic impacts of diarrheal disease in the cattle industry are also significant. Increased operating expenses are observed for dairy farms, cattle rendering plants and other livestock and poultry businesses that must increase su rveillance and management measures to comply with increased federal regulatory mandates (CDC 2002). Courts can impose stiff fines upon many different industries including producers, distributors, and restaurants.

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67 Escherichia coli O157:H7 Pathogenicity Esc herichia coli O157:H7 possesses multiple virulence factors, including production of Shiga toxins (Stx1 and Stx2) (Renter et al. 2004 a ), also termed verotoxins (Kruger et al. 2007) Escherichia coli O157:H7 is closely related to other STEC organisms, including Shigella species, as well as other strains of E. coli such as E. coli O55:H7, O111:H8 and O26:H11 (Renter et al. 2004a ). Of the Shiga toxinproducing E. coli strains, E. coli O157 is isolated from humans most often, and is responsible for most cases of HUS (Wang and R eeves 1998). Tarr et al. (1995) reported that E. coli O157:H7 was responsible for approximately twothirds of HUS cases in North America and Europe. The second most common E. coli serotype that causes HUS are E. coli O111 in the United States and E. coli O 26 in Europe (Monday et al. 2007). In Europe, a strain of E. coli O157 (SFO157) that, unlike E. coli O157:H7, does not ferment sorbitol, has been implicated as a cause of HUS (Monday et al. 2007). Infections of E. coli O157:H7 can lead to severe symptoms i ncluding hemorrhagic colitis (HC), and acute kidney failure due to hemolytic uremic syndrome (HUS) (Ogden et al. 2001). Hemolytic uremic syndrome is the most common cause of acute kidney failure in North America (Karmali 1989). In the United States, HUS is the primary cause of kidney failure in children (Breuer et al. 2001). Since it was initially identified in 1982 (Riley et al. 1983), E. coli O157:H7 has caused enteric outbreaks with 8% (Slutsker et al. 1998) and 9% (Bell et al. 1994, CDC 2009) of cases p rogressing to HUS (Slutsker et al. 1998). Progression of E. coli O157:H7 infections to HUS occurs throughout the world, especially in developed countries (Ooka et al. 2009). In Japan, 6% of persons infected with E. coli O157:H7 developed HUS (Watanabe et a l. 1999). In Scotland, E. coli O157:H7 has been determined responsible for more than 90% of HUS cases (Pollack 2005).

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68 In the United States, infection with E. coli O157:H7 was made a nationally notifiable disease in 1995 (CSTE 2005). Enterohemorrhagic E. co li O157:H7 (EHEC) is commonly found in association with livestock, especially cattle (Rasmussen et al. 1999, Hussein and Sakuma 2005). In the United States, bacterial outbreaks due to E. coli O157:H7 have occurred with increasing frequency since the discov ery of this pathogenic strain (Hancock et al. 1994). Escherichia coli O157:H7 P revalence and P ersistence Escherichia coli O157:H7 can persist in a wide variety of substrates for varying lengths of time. These substrates include livestock manure, human was te, garbage, compost, soil, and food crop plants, animal watering troughs, and natural water sources such as lakes, rivers and puddles (Avery et al. 2008). Persistence of E. coli in livestock fecal matter has been observed in manure from dairy cattle, beef cattle, swine, and sheep. Escherichia coli O157:H7 has also been observed to persist for up to three days in the acidic gut and feces of the house fly (Kobayashi et al. 1999) Escherichia coli O157:H7 can survive for up to 2 yr in dairy farm environments (Shere et al. 1998). This increases its potential for disease transmission. Longitudinal prevalence (i.e., persistence over long period of time) of E. coli on dairy farms is seasonal, with highest prevalence in spring and late summer (Hancock et al. 1997, shere et al. 1998, Vidovic and Korber 2006) Persistence of E. coli O157:H 7 has been observed in soil (Islam et al. 2004), water (Sargeant et al. 2003) and for up to 21 mo in composting manure (Kudva et al. 1998). Islam et al. (2004) observed that E. coli O157:H7 can survive in soils amended with cattle manure for more than five months. Persistence of E. coli O157 and specifically, E. coli O157:H7, in farm environments has been linked to its prevalence in water, including water tanks (Sargeant et al. 2003) and open water (Shere et al. 1998). Sargeant et al. (2003) reported higher individual cattle shedding rates

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69 for beef cattle in feedlot pens that tested positive for E. coli O157 than for beef cattle in pens that test ed negative for E. coli O157. They also observed that E. coli O157 was more likely to infect the water if it was detected in water tank sediment. Our understanding of the ability of E. coli O157:H7 to persist on plants has been further understood by recen t evidence that plants take up the bacterium through their stomata and translocate the organism to their tissues (Teplitski et al. 2009) However, isolation frequency of E. coli O157:H7 serotype has been lower than that of other enteric pathogens (Tyler and Triplett 2008) The ability to find "safe harborage" inside plant tissues could potentially permit greater persistence in plants than is possible by mere surface contamination. Persistence of E. coli O157:H7 and other enteric pathogens in and on plants is closely associated to the common agricultural application of animal manures (Brandl 2006) Escherichia coli O157:H7 Detection, Isolation and Identification Several different selective media have been used to detect, isolate and identify E. coli O57:H7 in samples collected from dairy cattle and other livestock farm environments, including Sorbitol MacConkey (SMAC) agar, Levines Eosine Methylene Blue (L EMB) agar, and CHROMAgar. These media contain selective agents, such as dyes and bile salts that inhib it the growth of competing microbial organisms, such as gram positive bacteria. Their specificity is increased further by addition of antibiotics (Heuvelink 2002). Although E. coli O57:H7 is difficult to isolate from fecal samples, it occurs in low densities as compared to other enteric microorganisms. In fecal samples, enrichment culture provides more sensitive isolation of E. coli O157:H7 than direct culture (Sanderson et al. 1995, Zhao et al. 1995, Wallace and Jones 1996), largely due to the inclusion of selective agents and/or antibiotics. Although selective media inhibit competing bacterial growth to improve recovery of E. coli O157:H7, each selective medium used to isolate and identify E. coli O157:H7 has

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70 disadvantages. For example, although SMAC agar contains selective agents to increase its specificity for isolation and identification of E. coli O157:H7, SMAC agar has poor overall specificity. Another disadvantage of SMAC agar is its propensity to change colors when incubated for prolonged periods, ma king it difficult to interpret results. Although supplementation of microbiological media, both selective and nonselective (general growth) types, with cefixime and potassium tellurite increases specificity, such supplementation can result in false positi ves, particularly when genetically similar enteric bacteria such as P. mirabilis or E. hermanii (Wallace and Jones 1996) are competing for growth. In addition to providing false negative results, supplementation with antibiotics and/or bile salts can actually inhibit recovery of acid or freeze stressed E. coli O157:H7, regardless of whether the media used is selective or non selective (Stephens and Joynson 1998). Escherichia coli is commonly found in complex media such as manure (Pell 1997) Although widely present in fecal matter, E. coli O157:H7 is difficult to isolate, because there are also high concentrations of competing, "background" microorganisms (BM), such as Proteus, Klebsiella, Salmonella, and Shigella (Vold et al. 2000) Escherichia coli populations decrease rapidly after host excretio n, because of changes in temperature, light, pH and moisture conditions (Unc and Goss 2006) and from increased growth rates of competing bacteria. Historically, the largest concentrations of E. coli have been obtained from fresh manure, either during or i mmediately after deposition.(Bolton et al. 1999, Duffy 2003). More recently, still higher numbers of E. coli have been collected by insertion of anal rectal swabs (Chapman et al. 1997, Pearce et al. 2004, Ahmad et al. 2007), a sampling method that is consi dered by some to be more sensitive than collection of deposited manure (Rice et al. 2003) However, Vidovic and Korber

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71 (2006) determined that large 10g fecal samples provided more accurate isolation of E. coli than swabs. Isolation of E. coli strains from manure is complicated by the large numbers and variety of different bacteria species present. Kudva et al. (1998) and Tutenal et al. (2003) observed that animals inoculated with 105 CFU/g of E. coli had a verage initial E. coli concentrations of 105 108 CFU/g in fresh feces, that concentrations in manure piles remained at 105 106 CFU/g for approximately one year, but decreased to 101 102 CFU/g when aerated (Tutenel et al. 2003, Dunn et al. 2004b) dete rmined that prevalence rates of E. coli O157 in dairy cattle manure were commonly < 1%, and never exceeded 5%. Pearce et al. (2004) tested up to three samples from individual cow pats, and determined that E. coli O157 distribution within single cow pats i s highly variable; thus, prevalence rates can vary significantly, depending on where in the cow pat samples are obtained. Echeverry et al. (2005) also tested multiple samples per cow pat, and determined that prevalence rates increased from 8.2% when one sample/pat was tested, and up to 20% when 5 samples/pat were tested. Isolation of E coli strains differs for diarrheagenic versus non diarrheagenic species (Heuvelink 2003) Within diarrheagenic E. coli strains, differentiation of enterohemorrhagic E. coli (EHEC) O157:H7 can be accomplished by any of several methods, and there is no general consensus on which me thod is superior. Typical E. coli biochemical characteristics include the following: Lysine +, Citrate Indole +, Acetate +, Lactose +, and aerogenic + (production of gas with carbohydrate metabolism). Escherichia coli O157:H7 does not ferment sorbitol within 24 h (Doyle and Schoeni 1984) and is therefore described as "non sorbitol fermenting" (Desmarchelier et al. 1998) Because most isolation culture methods involve 24h incubation, this characteristic provides easy

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72 differentiation from sim ilar and closely related enteric bacteria species. Escherichia coli O157:H7 has a characteristic morphology and color when grown on Sorbitol MacConkey agar (SMAC) supplemented with selective antibiotics such as potassium tellurite and cefixime (CT SMAC) th at inhibit gram positive bacterial growth and enhance gram negative enteric bacterial growth. Such media is commonly used, although the percentages of the respective antibiotics may differ among researchers. Typically, antibiotic concentrations are: potassium tellurite (1.25 g/l), sometimes referred to simply as "tellurite," (Alam and Zurek 2004) and cefixime (15 g/l) (Desmarchelier et al. 1998) Antibiotics are added to provide selective growth of enteric bacteria, especially of E. coli O157:H7, a s well as to provide differentiation of E. coli O157:H7 from its close enteric relatives. When cefixime and potassium tellurite are supplemented into SMAC, the resulting agar is referred to variously as CT SMAC, SMACCT, or even as mSMAC, where the "m" rep resents "modified," even though the type of modification may differ dramatically. Comparison of results obtained from use of SMAC modified with antibiotics needs to be conducted very carefully, for upon examination, not all researchers use the same concent rations of antibiotics. Thus, while multiple researchers may all refer to "CT SMAC," they may be using fundamentally different selective media. Additionally, some researchers add yet another antibiotic, vancomycin; again, with differing concentrations. In some rare instances, the antibiotic cefsulodin has been used, either in addition to those listed previously, or as a replacement for one or more of them. Cefsulodin is recommended along with cefixime and potassium tellurite in an FDA protocol for isolation of E. coli O157:H7 (FDACFSAN 2007b) There is one caveat regarding the use of potassium tellurite antibiotics in media. Although primarily used to select for nonsorbito l fermenting E. coli O157:H7, potassium tellurite also permits growth of other Shiga toxin producing E. coli strains such as O26, O111,

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73 and O145 which possess the ter gene that confers resistance to this antibiotic. Possession of the ter gene appears to be positively correlated with possession of the eae gene. Therefore, use of this antibiotic in selective plating media makes isolation of O157:H7 more likely, but not guaranteed. Strains of E. coli isolated with potassium tellurite tend to possess the eae ge ne (Orth et al. 2007) Even without supplemental antibiotics, SMAC is selective for gram negative bacterial growth, because crystal violet is a selectiv e component of sorbitol MacConkey agar (SMAC) that inhibits growth of competing enteric gram positive bacteria (Doyle and Beuchat 2007) For this reason, SMAC is wi dely considered to be the best media to use for successful isolation of E. coli O157:H7 (CDC 1994) Chou et al. (2000) observed that E. coli O157:H7 cultured in nonselective general growth trypticase soy broth (TSB) culture tubes for one day at the low temperature stresses of 5 C, 18 C and 28 C exhibited respective survival rates of 87.55%, 0.72%, and 1.66%, when followed with a subsequent 1h exposure to crystal violet. This high mortality of cold stressed E. coli O157:H7 after 1 h exposure to crystal violet illustrates the importance of crystal violet dye for the s elective isolation of E. coli O157:H7, even without antibiotics. The use of antibiotics is helpful against competing gram negative bacteria after crystal violet inhibition of gram positive bacteria. Although it may seem frustrating that researchers use va rious concentrations and mixtures of antibiotics to supplement SMAC for selective growth and isolation of E. coli O157:H7, there is ample justification for it: different strains or serotypes of E. coli O157:H7 may react dissimilarly when exposed to environmental stresses. Development of antibiotic resistance in a given bacterial population may change an organisms ability to grow in antibiotic treated media. Escherichia coli O157:H7 is typically resistant to many antibiotics, and will usually grow

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74 on media that contains cefixime and potassium tellurite, whereas other enteric bacteria will be inhibited by these two antibiotics (Desmarchelier et al. 1998). While CT SMAC, in all its variations, appears to be the most selective and most effective media for diff erential isolation of E. coli O157:H7, direct plating may not be sensitive enough to identify the low levels of O157:H7 that are typical of samples with low concentrations of E. coli O157:H7 such as exist in many food samples (Willshaw et al. 1994). This m ight also be true for fecal samples, particularly because cattle shed E. coli O157 both seasonally and sporadically, so that bacterial concentrations may vary widely from one sampling occasion to the next (Matthews et al. 2005). This lack of sensitivity le d to the development and wide use of immunomagnetic separation (IMS) for selective isolation of E. coli O157:H7 on magnetic beads coated with anti E. coli O157:H7 antibodies. IMS permits very selective binding of E. coli O157:H7 to the beads. The conjugate d bacteria bead complex is manually separated from bacterial enrichment broths magnetically and by simultaneous rinses in buffered detergent, such as PBS Tween 20. The benefit of IMS is that it permits a more sensitive detection of E. coli O157:H7 than tra ditional direct plating (Grif et al 1998). However, the disadvantage of IMS is that specificity may be decreased by the many sorbitol nonfermenting microorganisms that bind nonspecifically to the immunomagnetic beads (Chapman and Siddons 1996). Standar d microbiological selectivity of isolation has been greatly improved by immunomagnetic separation (IMS) technology which increases sensitivity 100fold over direct culture methods (Karch et al. 1996). Immunomagnetic separation utilizes magnetic beads coated with polyclonal pathogenspecific antibodies into specimen enrichment samples for inoculation. The magnetic beads specifically bind to the O157 somatic antigen of E. coli O157. Sequential rinsing steps remove any sample debris and/or nonspecifically bou nd pathogens from the beads,

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75 and the beads are then plated directly onto selective media (Heuvelink 2003). The IMS protocol is especially effective in samples such as feces which contain large populations of competing (background) microorganisms (Manafi 2003). Detection of E. coli O157:H7 by direct culture is an expensive, difficult, labor intensive and time consuming process with little consistency in chosen isolation methods among researchers (Manafi 2003, Heuvelink 2003) Standard microbiological tests typically take 3 4 days to complete and require repetitive re plating of suspect organisms onto new media each day. High background microfauna concentrations typical of fecal samples greatly increase the difficulty with which E. coli O157:H7 is successfully isolated. Competing microorganisms can be so numerous that they result in lawns of bacterial overgrowth, even when plated onto selective media plates that contain antibiotics specifically designed to inhibit the growth of non E. coli O157:H7 organisms (Heuvelink 2003) Identification by bi ochemical and morphological traits can take as long as 7 d, depending on how many traits are examined and which selective media are used. Biochemical tests have low specificity and selectivity (Visetsripong et al. 2007), making it difficult to accurately identify and isolate O157:H7, especially if present in very low numbers. Successful detection and isolation of E. coli O157:H7 is further confounded by the presence of closely related bacteria species. This is particularly evident in fecal environments whe re competing microorganisms share similar phenotypical and biochemical traits with E. coli O157:H7, even when isolates are plated on selective media designed to differentiate between bacteria species. Thus, isolation of individual colonies with typical E. coli O157:H7 characteristics on any particular media require subsequent transfer to additional selective media to confirm identification.

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76 Rapid, selective and sensitive detection methods are important (Ogden et al. 2001). Although addition of inhibitors a nd/or antibiotics to growth media increases the selectivity of the media for successful isolation of the target bacteria, it often decreases the sensitivity, so detection of small numbers of bacteria becomes increasingly more difficult. Escherichia coli O 157:H7 and DNAbased Isolation Techniques Polymerase chain reaction (PCR) provides a more selective and sensitive method than directculture to identify E. coli O157:H7 in samples (Visetsripong et al. 2007), and permits serotyping (Beutin et al. 2007). PCR can be performed in 6 hours (Szalanski et al. 2004), making it more promising for timely identification of E. coli O157:H7. Primer pairs can be used individually in uniplex or combined in multiplex PCR to amplify one or more specific target gene fragment s, respectively. Shiga toxin (Stx) gene fragments are often targeted in PCR because they are probably the main virulence factors (VFs) of E. coli O15:H7 that lead to HUS and HC (Beutin et al. 2007, Krger et al 2007). Successful PCR amplification of Stx gene fragments confirms the presence of the Shiga toxinproducing gene, but does not specifically confirm E. coli O157:H7, because Stx genes are also present in more than 200 serotypes of E. coli (Beutin et al. 2007) and in closely related Shigella spp. (D onnenberg 2002). While PCR provides quick identification of positive E. coli O157:H7 samples, one disadvantage of using PCR is that fecal components inhibit polymerase chain reaction, and thus result in false negative PCR results (Wilde et al. 1990, van Z wet et al. 1994) However, these chemicals can be rem oved by detergents (Wilde et al. 1990). Another disadvantage of PCR is that there is evidence that E. coli O157:H7 strains can lose Shiga toxin producing genes (Feng et al. 2001) so that PCR might not detect positive E. coli O157:H7 samples that are detect ed by serological or biochemical methods.

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77 Summary While cattle are a primary reservoir of E. coli O157:H7, they shed this pathogen intermittently in their feces (Wetzel and LeJeune 2006) The highest cattle shedding of E. coli O157:H7 occurs during warm s ummer months when both E. coli O157:H7 and house fly populations are also at their highest levels (Alam and Zurek 2004) Standard fecal cultures based on 25 g of cattle feces inoculated in 225 ml of a nonselective nutrient broth typically detect E. coli O 157:H7 concentrations that range from 2 x 102 to 8.7 x 104 CFU/ml. (Hancock et al. 1997) In contrast, adult house flies weighing only 0.13 g have E. coli O157:H7 in concentrations ranging from 3 x 101 to 3.0 x 105 CFU/fly (Alam and Zurek 2004) the equi valent of 3 x 102 to 3.0 x 106 CFU/ml, so there is as much as a 100fold higher concentration of E. coli O157:H7 in house flies than in manure. Thus, it is possible that E. coli O157:H7 on dairy farms might be more accurately detected by testing adult house flies instead of cattle manure samples, regardless of which isolation method is utilized.

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78 CHAPTER 2 HOUSE FLY DISPERSAL Introduction Musca domestica L., the house fly, is capable of transmitting more than 100 pathogenic organisms (Greenberg 1973) that can cause disease in humans. Musca domestica has especially strong links to enteric diseases, such as typhoid, cholera, dysentery, and diarrhea (Steinhaus 1946), which cause high mortality rates worldwide, particularly in children of developing countrie s (Kosek et al. 2003, iOWH 2008) The house fly's ability to transmit path ogens is due to the following synergistic factors: 1) its predilection to breed in fecal or rotting organic material that may be teeming with disease causing microorganisms, 2) its habit of constant regurgitation and excretion while eating, and 3) its abil ity to disperse over wide geographic areas as far as 33 km (20 mi) (Murvosh and Thaggard 1966, Meerburg et al. 2007) including direct flight over large swamps and across rivers 300 500 m wide (Shura Bura et al. 1962). Considered alone, none of these behaviors creates a human disease threat. However, cons idered together, the potential for disease transmission expands exponentially, as the house fly can potentially transmit many fecal pathogens to any food source within its dispersal range. Dispersal of the pathogeninfected house fly from a dairy to a town introduces and increases the potential for disease transmission to humans. Flight and dispersal behavior for house flies and other species of synanthropic flies with similar breeding habits, such as stable flies and blow flies, has been reported by a num ber of researchers and varies drastically in individual studies. In rural areas, house flies can disperse 12 km (Broce 1993 a ) and have been documented dispersing up to 21 km (Bishopp and Laake 1921, Alam and Zurek 2004) from their breeding sites. In urban

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79 communities, most flies disperse within 1.7 km (1 mi) of release sites (West 1 951, Quarterman et al. 1954, Schoof and Siverly 1954b; Hanec 1956; Sacc 1964, Milio et al. 1988) However, house flies have been documented to disperse distances up to 33 km in urban environments (Murvosh and Thaggard 1966). House fly dispersal speed has been documented at a rate of 1 km/h for the first 34 h, when dispersal occurred as direct flight over a large swampy area and acro ss rivers (Shura Bura et al. 1962). Dispersal distances and recapture rates might be influenced by the type of flies used, i.e., fieldcollected or laboratory reared. Previous studies have indicated that use of field collected flies is more representative of dispersal under natural conditions than flies that are reared for multiple generations in the laboratory. Eddy et al. (1962) recaptured a 10fold higher percentage of field collected flies than laboratory reared flies, implying that laboratory colonies may lose their ability to disperse. Dispersal of house flies increases the potential for transmission of zoonotic diseases to humans, particularly from sites conducive to fly breeding such as dairies that house large numbers of cattle (Kaufman et al. 2005 b, Ahmad et al. 2007, Conn et al. 2007) beef cattle feedlots (Skoda et al. 1993, Thomas 1993, Sanderson et al. 2006), swine facilities (Rosef and Kapperud 1983, Halverson 2000) poultry buildings (Hald et al. 2004, Watson et al. 2007) as well as non agricultural sources of fecal waste, such as dog excrement (Wilton 1963), to nearby human population centers. If house flies can maintain a travel speed of 1 km/h for an extended period of time, and if h ouse fly dispersal flight occurs in a straight line from a breeding site, then house flies could potentially transmit infectious pathogens as far as 12 km in only 12 h (Meerburg et al. 2007).

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80 The house fly readily alights on decomposing fecal/organic mat ter as well as human food, and moves freely between the two. External contamination of house flies can range from 2.5 to 29.5 million bacteria per fly (Hawley et al. 1951), and some bacteria can survive up to 3.5 d on the surface of house flies (Peppler 1944). Bacterial contamination of house flies can also occur after flies contact food crops that have been fertilized with liquid manure or solid fecal waste (Islam et al. 2005). Mechanical transmission of this bacterium by house flies has been well established by many researchers (Echeverria et al. 1983, Fotedar et al. 1992, Sasaki et al. 2000, Alam and Zurek 2004, Buma et al. 2004, Ahmad et al. 2007, Nmorsi et al. 2007). Biological transmission also appears likely if E. coli O157:H7 is capable of replicating within the house fly gut (Hawley et al. 1951, Petridis et al. 2006). House fly dispersal has been measured by many types of mark, release and recapture studies using fluorescent dusts, sticky traps, and UV lights (Hogsette 1983, Osek 2001). One of the easiest and most efficient techniques for marking and releasing of large numbers of small insects is the application of fluorescent dust (Hagler and Jackson 2001). Insects such as ho use flies are collected in the field or massreared in the laboratory, marked for future identification, released in the field, and recaptured at various distances from their release site. Ideally, the substance used to mark the insects is longlasting, is nontoxic to both the insect and the environment, does not change insect behavior, and is easy to observe on recaptured specimens. Additionally, the ideal insect marker is inexpensive, readily available, has a long shelf life, performs consistently, and c an be applied quickly and easily (Osek 2001). Quick and easy application of markers is

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81 particularly beneficial in the field, where access to tools used to decrease insect activity for easier application of markers, such as chill tables or CO2 anesthetic, i s limited. Many types of markers have been used effectively in the past, but are no longer recommended or are sometimes prohibited under existing regulatory legislature, due to human, animal and/or environmental health concerns. For example, radioactive phosphorus (32P) has been added to adult fly laboratory diets and 32P labeled flies released and recaptured in the field (Lindquist et al. 1951, Yates et al. 1952, Eddy 1962, Shura Bura et al. 1962, Williams, J. R. P. 1973). Marked flies were subsequently counted using readings on Geiger counters (Hoffman and Lindquist 1951; Lindquist et al. 1951; Yates et al. 1952; Eddy 1962; Shura Bura et al. 1962; Williams, J. R. P. 1973). Hoffman and Lindquist (1951) reared flies in media containing 32P to compare the efficacy of this method against application by ingestion, and deter mined that feeding 32P to adult flies was both more effective and cost effective. Lindquist et al. (1951) compared marking adult house flies by adding 32P to the diet against dusting with fluorescent dusts. They determined that marking with dietary 32P was more efficient and less labor intensive, and they observed that the dusts wore off within 48 h so that identification of dusted flies was difficult. Similarly, Eddy et al. (1962) concluded that ingestion of 32P was a more useful marking method than fluore scent dusts, because wild flies had natural fluorescence that was easily confused with the fluorescent marker (C 205 yellow, Ultra Violet Products, Inc.) used in their study. Although there is no universal method of marking insects, dusts are possibly the most frequently used external markers, due to their ease of use in both application and observation, as well as their low cost, ready availability, and low toxicity. Fluorescent

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82 dusts, in particular, DayGlo powdered pigment dusts, have been used to trac k dispersal and population dynamics, without any observed adverse changes to insect behavior or mortality (Hogsette 1983, Hogsette 1984, Kristiansen and Skovmand 1985). Fluorescent dusts also offer potentially longterm investigative study possibilities, because the dust has been shown to last up to 3.5 mo in the field (Taft and Agee 1962). Flies are dusted with fluorescent powder, released, and recaptured; subsequent UV light examination of recaptured flies illuminates any retained fluorescent dust on areas of the body that the fly has difficulty grooming. Flies dusted as adults will typically retain dust particles on portions of the thorax; when the puparia are dusted, flies emerge, crawl through the dust, and retain it on their ptinilum (Hogsette 1983). An additional advantage of fluorescent dusts is that their visibility is greatly enhanced when examined under long wave ultraviolet (UV) light. Thus, large numbers of recaptured house flies on sticky traps can be examined rapidly and easily under UV light to determine how many are marked. This eliminates time and labor intensive observation methods used with alternative marking techniques, as there is no need to destroy individual insects to observe internally expressed dyes, to apply solvents, or to perfo rm genetic analysis. Application of fluorescent dusts is relatively easy, inexpensive, and less labor intensive than other insect marking techniques, and enables marking of thousands of insects simultaneously (Zhao et al. 1999). Additionally, application of fluorescent dusts can be accomplished using mechanical dusters (Hogsette et al. 1993). Flies can be captured on a wide variety of traps. Alsynite traps have proven effective, easy to transport, and easy to use in the field. One type of alsynite trap consists of a translucent rectangular fiberglass panel folded to form a cylinder that is then

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83 wrapped with an adhesive coated clear plastic sleeve (Hogsette and Ruff 1990). The fiberglass panels reflect ultraviolet light, making them attractive to many flying insects such as stable flies, Stomoxys calcitrans (L.), (Williams, D. F. 1973), alate red imported fire ants, Solenopsis invicta Buren, (Milio et al. 1988), and house flies (Geden 2006). Flies that land on the adhesive become stuck. Traps can be examined under UV light to observe fluorescence on individual flies, and flies can be identified to species. Although flies ostensibly have all their physiological needs met in the dairy environment and have no discernible reason to leave the dairy, there is evide nce that they disperse in all directions without aid of vehicle transport, in direct flight, away from the dairy. The true pathogen transmission potential of M. domestica in north central Florida can be better estimated if house fly dispersal behavior can be determined. The goals of this study are: 1) determine if house flies disperse from a dairy to a town, 2) determine recovery rates of marked flies, and 3) evaluate possible deleterious impact of fluorescent dusts used to mark the flies. Materials and Met hods Laboratory Facilities and Rearing Unless otherwise stated, all laboratory studies and fly rearing were done in the fly rearing laboratory at the USDA ARS CMAVE in Gainesville, FL. Standard USDA rearing conditions for all fly stages in the fly rearing chambers were 26 2 C, 60 5% RH, 12:12 L:D. (Hogsette 1996). Any mention of fly larval growth medium refers to the Gainesville House Fly Diet (GHFD) (wheat bran 50%, alfalfa meal 30%, cracked corn 20%) (Hogsette 1992) unless another medium is specifically described. Larval growth medium was moistened with water in a 1:1 (v:v) diet:water ratio. Adult house flies were

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84 provided wit h food comprised of 6:6:1 powdered milk: granulated sugar: powdered egg yolk, and ad libitum access to water (Hogsette et al. 2002). During this study, dairycollected adult and immature house flies were frequently reared in the laboratory. Fly transport cages measured 30.5 x 30.5 x 30.5 cm (Model 1450B, Bioquip, Rancho Dominguez, CA) and were used to move adult dairycollected flies back and forth between the field and laboratory, and to release marked adult flies at the field site. Rearing cages were eit her: 46 x 46 x 46 cm (Model 1450C, Bioquip, Rancho Dominguez, CA), or 45 x 36.25 x 36.25 cm (USDA) (USDA ARS CMAVE, Gainesville, FL). Regardless of which cage was in use, adult house flies were maintained under standard USDA rearing conditions as previous ly described. Approximately 100 g of fresh GHFD was placed in each cage in 237 ml S tyrofoam deli cups (Model # 8SJ32, Dart Water Corp., Mason, MI). Approximately 400 ml of water was placed in 473ml clear plastic deli cups (Model #L2516, Newspring Packagi ng, Kearny, NJ), and covered with a single layer of foam packing pellets to prevent drowning. In adult rearing cages, fresh dry diet was added to cages every 3 d, and additional containers of water were added if water levels dropped to less than 25% or if dry diet or water showed mold growth. Transport cages were in use for only a few hours, so replenishment of dry diet and water was not necessary. Description of Study Area The study area used from 16 October 2008 to 4 December 2008 and from 14 May to 25 J une 2009 is located in north central Florida, in a geographical region dominated by small livestock farms, primarily dairies and poultry broiler farms. The area was relatively flat, with gentle rises (hillocks). Patchy habitats of scrub forest were intersp ersed

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85 throughout the area, with fields often separated by tree lines (wind breaks). Dairy pastures on the release farm were irrigated with water and fertilized with slurry (liquefied manure), both of which were applied with water sprayers of various types. The study area was the area between a small town and the dairy upon which the flies were released. This dairy is hereafter referred to as "Dairy A." There were two other dairies nearby: both were adjacent to Dairy A with shared property lines. "Dairy B" w as located W of Dairy A, and "Dairy C" was located S of Dairy A. The town was located entirely within 3.5 km SSW of Dairy A, so that traps placed at the closest edge of town relative to Dairy A were located approximately 2.5 km from the dairy, and the trap that was placed in town furthest from Dairy A (trap 12, at Restaurant D) was located 3.5 km from the Dairy A release site. House flies were collected from both Dairy A and Dairy B; however marked adult flies were released only from Dairy A. Dairy A milke d between 400500 cows and maintained 4 5 bulls at the beginning of the study. Approximately 50% of the cattle were sold during the summer of 2009, so the herd decreased to 200 300 by the end of the study. The cattle grazed on Bahia grass fields during the day, and came to the opensided barn twice daily to eat grain and be milked. Hay was provided in the fields to supplement grazing. The barn contained 2 3 large cement water troughs located midway between the south and north barn edges, and was equipped w ith misters that sprayed water over the cows, and large fans that increased air flow within the barn. The barn was located atop a small hillock, and consisted of a large cement floor that was slightly graded to enhance twicedaily rinsing of feces and urin e into a cement culvert that emptied into a nearby cement wastecontainment system. The floor was rinsed by flooding it with water that was stored in a large tank located at

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86 the eastern end of the barn at the point of highest elevation. After cattle feeding was completed, a release valve on the tank was opened, so that the entire contents of the water tank flooded the floor of the barn, and rinsed fecal waste into the culvert. The barn had two cement feeding troughs, one along each of the south and north lengths. The feed troughs were located at ground level, at the edge of the barn's roof drip line, so that the grain was largely protected from rainfall except during windy conditions. The troughs did not receive water spray from the barn misters. After rai n, large puddles formed next to the feeding troughs, and some runoff entered the open ends of the feeding troughs. Grain was poured into the troughs mechanically by a truck twice daily. Cattle entered the barn approximately 3060 min before the grain was delivered, and also were observed eating grain that remained in the troughs from previous feedings, even if it was slightly moist from rain or infested with fly larvae. The feeding troughs consistently served as house fly breeding sites during this study. I ntermittent applications of toxic fly bait containing imidacloprid (QuickBayt, Bayer, Shawnee Mission, KS) and insecticides containing permethrin both inside and outside the barn were used to suppress adult house fly populations. Additionally, permethrin impregnated ear tags were used for hor n fly, Haematobia irritans (L.), control. Dairy B was a dairy with 500600 milking cows, approximately 30 bull calves, and 78 bulls. The cattle spent the majority of their time in opensided barns, where feed was available ad libidum When in the barn, cattle were packed so tightly that they were pressed together. Cattle were milked twice daily. Cattle sometimes grazed in the grass fields, and supplemental hay was available in the fields. There were two barns, respectively placed north and south of the m ilking parlor, which were identical in

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87 structure. Each barn contained one large cement water trough located in a corner. Several large ceiling fans were evenly distributed throughout the barns. Water misters were also located throughout the barns. The barns were located on an inclined patch of land so that the north barn had the higher elevation. The cement barn floors had little or no grade, limiting runoff. Fecal and urine waste products accumulated rapidly, and were removed by spraying water with a handheld hose. Liquefied waste flowed into a nearby earthen lagoon system situated south and downhill of the south barn. A large cement feed trough ran east to west through the center of each barn. Feed was provided daily to the trough by a mechanical feed auger system. The feed troughs were elevated, so that cattle could consume feed without stooping. Fecal and urine waste did not appear to contact grains. New feed was apparently added to old feed as troughs were never completely empty. The troughs were withi n range of the water misters, and the feed was frequently damp. The feed troughs were surrounded with steel poles that provided abundant horizontal and vertical resting surfaces for adult house flies. Additionally, rough wood support beams were located at regular intervals along the feed troughs. The barn's exterior was constructed of rough wood, with wood beams extending from floor to ceiling, and plywood panels covering the top half of the barn. Cattle were restrained in the barn by a wood fence that surr ounded the perimeter of the barn. The south barn was built so that its southern edge was above ground. Fecal waste draining from the south barn usually overflowed onto the ground, and possibly contaminated spilled grains that were located underneath the a uger used to supply new feed to the feed trough. Overflow of fecal waste into spilled grains was further facilitated by a constantly flowing hose that supplied the water trough inside the barn. The trough

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88 overflowed out of the south barn onto the ground through which the fecal waste moved en route to the lagoons. The spilled grains were located slightly downhill of the fecal waste ditch. Therefore, grains that spilled onto the cement pad beneath the auger were moistened by various sources, including rainfal l and diluted fecal waste. Between the south barn and the lagoons, there was a large patch of untended grass and herbaceous growth. Some of the calves were kept in individual calf hutches, while others were permitted free range in a field. This field was not mown during the 2009 6 wk study and the forage growth was taller than calf height by the end of the study. The calves were all located in a pasture approximately 0.5 km south of the feed barns. Calves in hutches were supplied with feed and water. Hutc hes were placed in the shade under large deciduous trees. Feed was provided to the calves in a large plastic drum s with and without protective coverings. Water was provided by insertion of a water hose into a large plastic drum set on its end. Similar to Dairy A, fly control was attempted by scattering imidacloprid fly bait and spraying permethrin around barns. Additionally, permethrinimpregnated ear tags were attached to cattle ears. House Fly Collection and Rearing Adults and immature stages of house f lies were collected from both dairies weekly for this mark, release and recapture dispersal study. Adult house flies were collected via repeated sweep netting above the feed troughs for up to 30 min by 12 persons, and transferred into a transport cage tha t already contained water and adult house fly diet, as previously described. Transfer of flies from sweep nets into transport cages was accomplished by inverting the net into the cage through a cotton sleeve

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89 attached to the front of the cage, and shaking f lies loose inside the cage. Adult flies that were captured and subsequently released on site at Dairy A in this manner are referred to as the sweep netted flies (SN). The total number of adult house flies captured each week was estimated using the followi ng two methods. During the first week, an average sweep net count rate was established for each individual, by having each person collect two extra sweep net collections, one at the beginning of sweepnetting, and one at the end of sweepnetting. When dair ycollected house fly populations at the dairy were visually low, the number of flies caught in initial sweeps was much greater than the number caught in final sweeps. The first and final sweeps were used to determine average sweep net values per individua l researcher, to compensate for the decreased house fly population in the vicinity resulting from the sweep netting impact. After determining each researcher's average sweep count rate, the total number of house flies collected per individual was estimated by multiplying the number of sweeps that each person performed by that individual's respective average sweep net count rate. Finally, the total count of house flies captured for the mark and release technique was calculated by adding individual counts together. This method was used only during the first week, because fly numbers appeared to be reduced too much by this method. Therefore, during the remainder of the study period, visual estimates of the reared adult house flies were made. To supplement dair ycollected caught adult flies, immature dairy collected flies were reared in the laboratory (IR). House fly eggs, larvae, and/or pupae were collected by filling 3.79 liter plastic buckets (Model # 86712 NRC 65 MIL, Letica Corporation, Rochester, MI) with decomposing grains and other dairy sources (dairycollected

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90 materials) where immature flies were observed prior to collection. Dairy collected materials and immature flies were transported back to the USDA laboratory in an air conditioned vehicle. In the laboratory, dairycollected materials containing IR developing flies of all immature stages were evenly dispersed among several enamel coated steel dental pans (19 x 31 x 5 cm) so that each pan was filled to a depth of approximately 2.5 cm. The remaining depth of each pan was filled to the top by addition of moistened Gainesville house fly diet (GHFD) (Hogsette 1992), as previously described. Larval rearing pans were placed individually inside Bioquip rearing cages. Adults were permitted to emerge from the rearing material, and were provided with adult house fly diet and water as described previously, in the rearing cages, for up to 6 d. After 6 d, all emerged adult flies were transferred to a transport cage as described below. In preparation for transport to the field, adult IR house flies were transferred from the rearing cages to smaller transport cages using CO2 introduced to a cage placed in a plastic bag. Flies were examined after each application of CO2 for recovery. Anaesthetized adult flies were gently shaken out of the rearing cage and into a transport cage. Flies were provided water and adult house fly diet during transport back to the dairy. The IR adult flies thus produced from dairycollected caught larvae ranged in age from 1 6 d (some adults emerged within hours of collection), depending on the day of emergence during the week spent in the rearing cage. After adult flies were removed from the rearing cages and transferred into the transport cages, the adult house fly diet, water, and larval r earing pans containing dairycollected larvae and pupae were placed back into the rearing cages to allow the

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91 emergence of additional IR adult house flies. These rearing pans continued to produce adult house flies for up to 3 wk. To further supplement numbers of adult flies, gravid dairycollected females were used to produce an F1 generation of adults that were subsequently marked and released at the dairy. Parental adult house flies were collected in the field by sweep netting, and transported back to the USDA fly rearing laboratory, as described previously. Oviposition chambers were prepared for each rearing cage and consisted of approximately 400 ml of 1:1 (v:v) water:GHFD (Hogsette 1992) as described above, placed in a 473ml clear plastic deli cup (Mod el #L2516, Newspring Packaging, Kearny, NJ). Oviposition chambers were placed inside each rearing cage for up to 3 d to provide oviposition sites for dairycollected house flies. After 3 d, the moistened GHFD containing F1 dairy collected house fly immat ures was removed from each oviposition chamber and added to a fresh mixture of moistened house fly diet in a large 58 x 46 x 8 cm rearing tray (Model #4003N, Del Tec Panel Controls, Greenville, SC) by spreading it evenly across the surface of the fresh di et. This facilitated movement of fly larvae into the fresh material, eliminating the need to separate larvae from spent material. The rearing trays were then placed inside a dark colored close woven cloth bag, which was twisted shut tightly and secured wit h a rubber band to prevent egress of adult house flies. These covered trays containing the F1 larvae were placed in the previously described rearing room, and were examined daily until pupation occurred. After pupation occurred, the pupae were separated f rom the diet via flotation, then gently air dried and separated from chaff in a forced air chamber (Bailey 1970, Hogsette 1992) for up to 2 h. Finally, the clean, laboratory produced F1 house fly pupae were

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92 permitted to emerge in a transport cage which alr eady contained 1to 6d old IR adult house flies that had emerged from dairy collected collected larvae. Because oviposition chambers were left in the rearing cages for up to 3 d, F1 adults emerged over a 3d period. Because daily emergence data were not recorded, the percentages of flies for each daily age are unknown. On the next scheduled field release day, cages containing all adult IR and F1 house flies were transported to the release point. The age and sex of flies that were marked and released wer e not recorded. Transport of Adult Flies to the Field During transport to the field, food and water were provided ad libidum as described previously. The transport cage containing adult flies was placed within a 41 x 42 x 35 cm cardboard box (transport box) and covered with a loose fitting lid. This box provided shade and was positioned in the center of the vehicle to further decrease sunlight impact upon the house flies. Marking, Releasing and Recapturing Adult House Flies Upon arrival at the release sit e, the transport cage was removed from the box, and placed in a sheltered, well ventilated, and shady location. The transport cage was stored in this protected location for 2 3 h while alsynite traps were placed in the field as described below and while additional adult house flies were collected by sweep net. After all alsynite traps had been placed, adult house flies collected by sweep netting were added to the transport cage and flies were marked with fluorescent dust in the following manner Corona mag enta (CM) and arcyellow (AY) Day Glo fluorescent dust (DayGlo Color Corp., Cleveland, OH) were applied on alternating weeks to the caged flies using

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93 metal plunger dusters (Hudson Manufacturing Co., Chicago, IL) to gently pump dust through the screen. Pri or to dusting, the cage was placed inside a large 170 L plastic bag to simultaneously enhance the application of dust to the flies and reduce the deposition of dust into the dairy environment. No attempt was made to quantify the amount of dust applied, or to calculate the amount of dust per fly. Dust was applied until it was visually apparent that all flies in the cage were well coated. After the dust was applied, the plastic bag was tightly closed by twisting and knotting the top. The bagged cage was gentl y shaken for 510 sec to disperse the dust onto caged house flies. The bagged cage was then set aside in the shade for up to 2 min to permit the dust to settle inside. After the dust settled for 2 min, the large bag containing the dusted cage with marked flies was opened carefully, and downwind of all personnel and vehicles, to reduce excess transfer of dust to the environment, and to prevent airborne dust from drifting onto traps without said transfer being performed by active fly transport. The lid of th e cage was lifted slowly and completely to permit egress of dusted adult house flies. Flies that remained in the cage after 1 min were removed by forcibly tapping the cage while holding it upside down directly above the plastic bag. This action resulted in a secondary dusting for these flies, as they landed in a pile of dust inside the plastic bag. This pile of dusted flies and excess dust was left undisturbed until all flies had groomed adequately to permit them to disperse out of the bag and into the dair y environment. Dispersal of all flies away from the dust pile was completed within 30 min. Commercially available alsynite traps (Olson Products Inc., Medina, OH) ( Fig. 21) were placed at approximately 0.5 km intervals between the dairy release point and the adjacent town. Traps consisted of 66 x 33 cm corrugated alsynite panels that were formed

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94 into a 30 cm diameter cylinder. Traps were secured into their cylindrical form with a 2.5 5.0 cm overlap by insertion of 2pronged metal fasteners into holes dril led along the edge of the panels. Traps were inserted into pre cut slits in either 2 x 2 x 50 cm wooden stakes (short stakes, provided with trap) or handcrafted 3 x 3 x 90 cm wooden stakes (long stakes). Regardless of size, stakes were inserted into the soil until the base of the trap was approximately 30 cm above ground, to provide a total trap plus stake height of approximately 65 cm. Sticky Sleeves (Olson Products Inc., Medina, OH), clear plastic sheets coated with an adhesive and protected with a waxy white peelable paper backing, were wrapped around the exterior cylindrical portion of the traps, and secured with 24 large paper clips. The waxy paper backing was then removed, and the sticky surface was exposed. Sleeves were labeled with trap number and date of placement on the nonsticky surface of the clear sheet using a permanent ink black marker prior to wrapping them around the alsynite cylinders. Some traps were placed along the edge of main roads between the dairy and town. Traps were positioned so that some traps could be used to determine corridor and habitat edge movement (Anderson and Danielson 1997, Fried et al. 2005) by placement along well travelled roads. Other traps were placed in patchy habitats away from roads. On the dairies, traps were placed near barns, milking parlors, and calf hutches. In town, traps were placed close to dumpsters outside two restaurants (traps 12 and 15), two convenience food stores (traps 17 and 23) and the local post office (trap 16), representing a nonfood site. Four traps (13, 14, 17, and 19) were moved due to difficult access or human animal disturbance after wk 2 and renamed (21, 22, 23, 24) s o that they were site specific. Geographical Information System (GIS) coordinates were recorded at each trap

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95 location and used to generate a diagram of trap placement, and trap locations were mapped on an aerial image (Google Earth 2009) ( Fig. 22). Direct distances to each trap from the release sit e were determined using the GIS data, and concentric arcs were used to roughly indicate 0.5 km direct distances. Trapping sites located between the 2.5 and 3.5km rings from the release site were within the town's border and traps at 3.5 km were at the far edge of town. In all instances, attempts were made to ensure that the traps were located as closely to the 0.5km concentric arcs as possible. In instances where such placement involved placing traps on nonpublic sites, permission was obtained by the respective homeowner or local business or public school. The purpose of placing traps in multiple environments allowed for both corridor movement and straight line flight assessment and enhanced the probability of recapturing dairyreleased, marked house f lies. Although it would be interesting to make the distinction between dispersal due to fly direct flight versus fly transport on automobiles from the dairy into the town, the data being sought here were solely to determine if dispersal was occurring rathe r than to determine how the dispersal was occurring. Sticky sleeves were collected weekly, and taken back to the laboratory for examination under a handheld 100W longwave UV light (Model # B 100AP, BLAK RAY, Upland, CA). Muscoid flies were identified to genus; house flies and stable flies were identified to species and the number of fluorescing house flies was recorded. Dispersal studies were performed from 16 October 2008 to 4 December 2008 and from 14 May 2009 to 25 June 2009. In 2008, 20 traps were p laced in the field weekly for 5 wk, for a total of 5 replications. In 2009, 18 traps were placed in the field weekly for 6

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96 weeks, for a total of 6 replications. Traps were placed in the field on Thursday mornings, and collected the following Thursday morni ng. House flies were marked and released within 1 h after placement of the last trap. Unless otherwise stated, traps were not visited during the 7 d. Trap maintenance was conducted so that new sticky sleeves were placed on traps immediately after the remo val of the 7d exposed sticky sleeves. Collection of used sticky sleeves was accomplished by placing a protective waxy paper sleeve cover on top of the adhesive side of the sticky sleeve so that captured flies were protected between sleeve and cover. This was secured in place with one large paper clip on each end, attaching it to a cardboard sheet that was cut to the same dimensions as that of the sticky sleeves. Precautions to minimize secondary transfer of fluorescent dust were taken. These included cove ring the sticky sleeves as described. Additionally, researchers involved in collection of traps used alcohol based hand wipes and/or 70% ethyl alcohol, to clean hands after processing each sticky sleeve. Both of these cleaning methods had proven successful in removing fluorescent dust from hands and surfaces in preliminary tests conducted prior to fieldwork. Finally, between field visits, the interior of the vehicle where traps were stored was wiped down with alcohol based wipes. Effects of Fluorescent Dust on House Fly Adults To evaluate the potentially deleterious impact of fluorescent dust upon dairycollected adult house flies, mortality was examined in the laboratory over 24 h under excessive dusting treatment cond itions (Fig. 2 3 ), hypothesizing that mortality due to excessive dusting would occur within the first 24 h. Adult house flies were collected using the aforementioned sweep net at the release site, placed in a transport cage, provided with food and water ad libidum as described previously, and transported back to

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97 the laboratory. In the laboratory, adult house flies were removed from the transport cage by anaesthetizing the flies with CO2, in a manner similar to that described previously to transfer flies from rearing to transport cages. The only difference in administration of CO2 was that the transport cage containing dairy collected house flies was placed inside a transport box and lid prior to being enclosed inside the large plastic bag. Because only a few flies were needed to test mortality i mpacts of dust treatments, three groups of approximately 100 anaesthetized flies were removed from the cage by gently scraping a small index card (62.5 x 75 mm) along the floor of the cage to scoop up the immobile house flies. Each group was placed in a cardboard ice cream cup (237 ml, Solo Cup Co., Urbana, IL) modified by removing the cardboard base and replacing it with window screen (940 x 813 microns (16 x 18 mesh); lids were placed upon the ice cream cup before flies revived. The experimental design consisted of three groups of 50 dairycollected house flies. Two treatment groups were marked with fluorescent dust (application technique described below): one group with 0.1 g of corona magenta dust, and the second group with 0.1 g of arc yellow dust. The third group served as a control and was not dusted; however, the ice cream cup "cage" was shaken in the same manner as if dust had been added. Dust was administered by placing the dust on top of the window screen, and gently pressing it through the windo w screen with a spoon. A second lid was placed over the screened base, and the cup plus the two lids were placed inside a self locking plastic bag (0.95L, Ziploc Racine, Wisconsin), and agitated for 510 sec. Dust was allowed to settle inside the cardbo ard cups for 515 min before opening the bags. Afterwards, the

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98 cups were removed from the bags, and the second lid was removed to reveal the screen base. Each cup was placed in a plastic deli container with lid, and CO2 was gently administered without dist urbing the dust to achieve knockdown. The flies were then placed on a sorting tray, where a low level release of CO2 kept the flies from egressing, but permitted leg twitching, while flies were counted into test chambers. Rectangular pieces of window screen (25 x 20 cm) were folded to form rectangular test chambers (12.5 cm x 20.5 cm). Test chambers were stapled along two edges, leaving one short edge open. Ten marked house flies were counted into each test chamber, and a cotton ball saturated with 5%sucr ose solution was placed inside to provide food and water to the flies and to prop the sides of the test chamber apart to permit fly movement within a 3 dimensional space. Test chambers were maintained in a laboratory at 3335% RH, 2628 C. House fly morta lity was assessed 24 h after dust treatment and flies that were ataxic were considered dead. Weather Weather conditions that were observed during the time spent in the study area were recorded on the days that fieldwork was conducted. To obtain more compl ete weather data for the entire study period, weather conditions were obtained from Weather Underground (2009) at a field recording station located 29 km NE of Dairy A. Downloaded weather data included daily temperature, relative humidity, precipitation, barometric pressure, wind speed, and wind direction. Daily data were used to generate weekly mean values for each climatic factor for the 7 d test period. The 7 d period for each week began on Friday, and concluded on the following Thursday when traps were collected, so that weekly mean values were for the 6 d prior to and including the day of trap collection.

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99 In association with extreme weather conditions, such as approaching storm fronts, adult fly behavior anomalies were observed. These behavioral anomal ies were recorded, particularly if they appeared to impact house fly dispersal. Additionally, any observations of unusual larval conditions, such as high mortality due to flooded larval breeding sites following heavy rains, were also recorded. Statistical Analysis Mean, minimum and maximum dispersal distances were calculated using Microsoft Excel (Excel 2002). Released and recaptured fly data were subjected to PROC UNIVARIATE to examine normality and PROC MEANS to calculate means using SAS Version 9.1 (SAS 2002). Paired correlations between numbers of re leased and recaptured flies were analyzed using PROC CORR (Pearson's coefficient) (SAS 2002) for both years combined, for 2008, and for 2009. These data were then submitted to linear regression analysis with recaptured numbered of house flies regressed against released numbers of house flies using PROC REG (SAS 2002). These data were regressed for both years combined, and for each year individually. Results The mean dispersal distance for marked flies each w eek was 0.22 km (range 0.00 0.35 km) in 2008 and 0.62 km (range 0.03 1.12 km) in 2009 (Table 2 1). The maximum weekly dispersal distance ranged from 0.001.00 km in 2008 and from 0.103.00 km in 2009. Two marked house flies were recovered in the nearby town in 2009 (traps 11 and 15) (Table 2 2). The estimated number of flies that were marked and released each week ranged from 5003,700 in 2008 and from 2,00010,000 in 2009 (Table 2 3). No flies were marked and released on 7 November 2008 due to a lack of SN collected flies. No flies were marked and released on 21 May 2009 due to severe thunderstorms and heavy rain

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100 that prevented collection of house flies. Approximately 9,200 and 28,000 house flies were marked and released during the 5 wk 2008 test and the 6wk 2009 test, respectively (Table 2 3). In 2008, 20 traps collected a total of 13,141 marked and unmarked house flies; of those, 106 were marked (Table 23). The 106 marked flies represented 1.15% of the 9,200 marked flies released during the 2008 test period (Table 2 3). In 2009, 18 traps collected a total of 48,435 marked and unmarked house flies; of those, 144 were marked (Table 2 3). The 144 marked flies represented 0.51% of the 28,000 marked flies released during the 2009 test period (Table 2 3). We ekly recapture percentages for marked flies ranged from 0.00 to 1.93% in the 2008 test period and from 0.12 to 0.81% during the 2009 test period (Table 23). Weekly numbers of marked flies that were released varied between 0 in weeks 4 and 7 and 10,000 in week 11. The lowest recapture rate of 0.00% in 2008 followed three weeks of cold air temperatures that ranged from a weekly average of 7.4 C to a weekly average of 12.3 C. The preceding t wo capture weeks also had lower recapture rates as compared to the recapture following the first two releases. Notably, f lies recaptured at the end of week 4 were part of a release of 500 flies in week 3, so that the 0.20% recapture rate of week 4 reflects flies that were in the environment in 2008 for 7 14 d (Table 2 3) The lowest recapture rate of 0.12% in 2009 was obtained at the end of week 7 when no marked flies were released. These marked flies that were captured on 28 May 2009 were actually part of a 6,000fly release made at the beginning of the previous week (we ek 6) and had remained in the local environment for 714 d (Table 23). The highest weekly recapture rate of 1.93% was obtained in week 2, following release of 3,000 marked flies.

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101 When the cumulative recovery of marked flies over distance from the release site was calculated, 99.1% (105/106) of the marked flies that had been recaptured were collected from traps within 0.5 km (<0.5 mi) of the release site in 2008. In 2009, 88.9% (126/144) and 93.8% (128/144) of the marked flies that had been recaptured were collected from traps within 0.5 km (<0.5 mi) and 1.5 km (<1.0 mi) of the release site, respectively (Table 2 2). Total, daily mean and trap mean numbers of all captured house flies including recaptured marked house flies are reported for each trap in Tabl e 2 4. Weekly mean weather paramet er s are shown in Table 24. There did not appear to be any relationship between house fly recapture rates and the weekly mean temperature, the weekly mean precipitation, or the w eekly mean barometric pressure. Dust treatments did not appear to impact dispersal, as m ortality following exposure to AY and CM dust was 4%, while 2% of control flies died over the 24h evaluation period. Production of adult house flies was increased by allowing dairycollected adult house flies to oviposit in the house fly diet, resulting in production of approximately 7,000 F1 adults in 2009. Of these, 4,000 were marked and released in wk 3 and 3,000 in wk 4, representing 40% and 50% of these releases, respectively. F1 progeny adults were simila rly aged (within 3 d) house flies. No F1 progeny were produced in 2008. The overall number of flies that were released was not correlated to the number of marked flies recaptured (r=0.72099, p=0.0123). When tested by year, there was still no correlation b etween release and recapture rates for either 2008 or 2009. Linear regression analysis results also indicated no correlation in 2008. However, a positive correlation existed in 2009 (df=1,4; F=9.28, P =0.0381). Multiple regression analysis for both years co mbined showed an overall positive correlation (df=1,9; F=9.74, P =0.0123). The linear

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102 regression line for both years combined is: recaptured_flies = 0.402 + 0.0066*released_flies. Discussion Insects disperse for many reasons, including: to avoid overcrowdi ng, to take advantage of extended resources available in nearby locations and, to avoid unfavorable conditions (Stein 1986). Dispersal of house flies and other insects of medical and veterinary importance needs to be more fully understood, because more tha n 50% of infectious diseases world wide are transmitted by insects (Stein 1986) Dispersal of house flies, and the impact of fly dispersal on transmission of human diseases such as epidemic diarrhea, has been scientifically observed for more than 100 years (Kumar and Carmichael 1998) Although the earliest studies are more anecdotal than quantitative, they nevertheless provide useful clues that have been confirmed repeatedly using quantitative techniques in the intervening century, regarding house fly behavior and environmental impacts upon dispersal. For example, Nash (1913) concluded after his 19041909 studies that house flies tend to remain within 0.8 km of their breeding site in more urban areas, but will readily travel 1.6 km in more rural locations with fewer human habitations. The tendency of house flies to typically disperse within only 0.81.6 km from a breeding or release site has been repeated consistently by multiple researchers working independently in different geographic locations, unique environmental conditions, and with different experimental designs (Quarterman et al. 1954, Lysyk and Axtell 1986, Schoof and Siverly 1954b, Nazni et al. 2005, Winpisinger et al. 2005). This is consistent with the current study, where the typical distance in 2008 was 0.5 km, and 1.5 km in 2009. For both years combined, >95% of flies were captured on traps placed less than 1.5 km from the farm.

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103 House flies can disperse over long distances in a short period of time. Peppler (1944) collected house flies that travelled 5 km in 1.5 d. Bis hopp and Laake (1921) observed that house fl ies travelled 8.3 10 km and as far as 21.3 km in 24 h. Shura Bura et al. (1956) reported dispersal of 34 km within one hour. Sacc (1964) reported house fly dispersal up to 8 km, while exceptional dispersal distances of >32 km and 33 km were reporte d by Schoof and Siverly (1954b) and Murvosh and Thaggard (1966), respectively. One of the three objectives of this study was to determine if house flies could be dispersing from the dairy to the town, a 3km straight line flight. Because one marked fly was recovered from a trap in the town, this hypothesis was confirmed by the data. In general, t his short distance is typical for house flies, whose flight range has repeatedly been reported to be within 1.6 km, despite the exc eptional distances travelled by individual house flies (Schoof and Siverly 1954a,b). Dispersal studies with house flies to determine the flight range have always been difficult to conduct successfully, especially in urban settings (Murvosh and Thaggard 1966). Urban communities offer many substrates that are attractive to house flies, including garbage, decaying grass, coffee grounds, and excrement from mammals and birds (Schoof et al. 1954b ). Schoof and Siverly (1954b) estimated that an individual house fly may travel 9 km in its lifetime, due to meandering movements between sites. In my study, traps were placed near garbage dumpsters outside two restaurants and two convenience stores in town, but not near residential garbage cans. If residential garbage is attractive to dispersing house flies in the current study area, then the dispersal data reported in my study would be under reported. In my study, dispersal of house flies was observed up to a maximum direct line distance of 3 km over a 7 d test period. H owever, because I collected traps only weekly, I

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104 was unable to determine dispersal rate. It would be informative to repeat this study with collection of sticky traps over shorter intervals, such as daily or hourly, to obtain detailed information about the temporal dispersal rates of house flies from dairies into this north central Florida town. It would also be useful to track fly meandering movement between sites by placing self marking traps (Hogsette 1983) with a unique color of dust at each site. By doi ng so, recaptured flies that possess more than one color of dust would clearly have been present at multiple sites between release and recapture. Although house flies may disperse away from a breeding site, their dispersal may be more complicated than mer ely traveling from point A to point B. There is evidence to substantiate observations that flies disperse with "randomized, reciprocal type meanderings" (Williams 1973) that result in back and forth travel between the breeding/release site and some other site(s) of interest as distant as 17 km (Schoof and Siverly 1954b). This type of dispersal is of particular importance in areas where large animal rearing facilities such as dairies with endemic pathogenic E. coli O157:H7 are located within close proximity to human populations because it could greatly increase the potential for pathogen transmission to humans. Because flies were only marked at the release site, I was unable to determine if captured house flies had been at an alternative attractant site other than the dairy release site. Future studies could examine the meandering behavior of house flies in this study area by release of differently colored marked flies at multiple locations within the study area. For example, the dusting of flies at several da iries and at the garbage dumpsters in town would allow observation of dispersal patterns within the entire study area.

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105 Dispersal of house flies may be complicated by behavioral tendencies of insects to use landscape features to facilitate movement along corridors or to follow habitatedge environments (Anderson and Danielson 1997, Fried et al. 2005, Reisen 2010) Furthermore, dista nces travelled by dispersing house flies are typically measured as direct flights from point A to point B, with no regard to potential fly dispersal along corridors such as highways and/or vehicle assisted movement (Nazni et al. 2005) If flies use highways as corridors for undirected dispersal movement, then distances travelled could actually be under representative of the capability of flies that travel directly to new sites. Interestingly, Quarterman et al. (1954) report that house flies have a living space of up to 6 km diam over which they roam freely. Flies travel randomly throughout this space and aggregate at multiple feeding and breeding sites so the populations at multiple sites becomes one huge metapopulation. Quarterman et al. (1954) also observed that house flies disperse rapidly away from the release site, and that trap catches were highest on the first 3 d following releases. Simila rly, Schoof and Siverly (1954b) demonstrated that house flies display random dispersal and directional reversals by using three secondary mark and release sites in addition to the primary release site. Their release of radio labeled flies from a primary si te followed by dusting and release and recapture of these radio labeled flies at three secondary sites indicated movement in all directions between the multiple sites. Interestingly, my study indicates that house flies dispersed by both direct overland f light and by corridor movement. Direct flight is strongly implicated by the recapture of many flies at a neighboring dairy located 1.5 km to the west, with no direct road access connecting the two dairies. These two dairies were separated only by a common fence

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106 line and patchy habitat that included tree wind breaks between their mostly open pastures that held cattle. Similar dispersal of house flies from one dairy to an adjacent dairy was also observed by Denholm et al. (1985) and Lysyk et al. (1986). Hous e fly movement along landscape corridors with barriers, such as tree lines, is also likely (Fried et al. 2005), and can occur by both vehicle assistance and by direct flight. Vehicle assistance is likely due to milk and feed trucks, and other service vehicles that visited both dairies on multiple occasions during the field study. Similarly, Sacc (1964), Nazni et al. (2005) and Sievart et al. (2006) reported passive transportation of flies between dairies by automobiles. H ouse flies were frequently observed on the exterior of the research vehicle after departing the release site, for up to 5 min. To reduce the potential for providing vehicle assisted transport to the flies, typical departure from the release site was followed by driving away from the town al ong roads for up to 15 min until no flies were observed on the vehicle. During this time, efforts were taken to rid the interior of the vehicle of any flies. Additionally, there was no travel in this study towards the town after releasing the marked flies. It is unknown what impact the milk and feed trucks, and other vehicles had on fly dispersal between t he release site and the town Using only traps located 0.5 3.5 km, and excluding the four traps that were removed from the study after wk2, the number and percentage of traps that were positive for recovery of a marked house fly were approximately equal for both roadside (5/8 traps, 62.5%) and patchyhabitat (7/11 traps, 63.6%) traps. Although the number of positive traps was very similar, the roadside traps recovered higher numbers (n=39) of marked flies than patch habitat traps (n=16). This suggests that flight by house flies along corridors (Fried et al. 2005, Reisen 2010) as reported for other insects, is more important

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107 than flight through patchyha bitats in this region of northcentral Florida. However, it also clearly indicates that both types of dispersal occurred, and that marked flies were disseminated throughout and active at multiple sites throughout the study area. Further dispersal by recapt ured individuals might have occurred had they not been prevented from continued travel by the adhesive sticky sleeve. One marked house fly was recovered at a trap placed at Restaurant C located at the maximum recapture distance (3.0 km) obtained in t his study. Although Restaurant C was located in a treecovered patchy habitat, it was also located close to the major road that ran east west through town. Therefore, no conclusions can be made regarding the probable dispersal method or direction of approach used by the fly that was recovered at Restaurant C (trap 1 5). The fly could have arrived there by undirected flight, visiting multiple attractive sites along the way (Schoof and Siverly 1954b), by following corridors provided by roads (Johnson 1966), or by direct flight through the woods flight (MacLeod and Donnelly 1960). Marked flies were recaptured every week except week 5 (Table 2 3), which followed several weeks of low er temperatures ( 7.2 to 9.5 C ) than surrounding weeks (Table 2 4). Dispersal in the study area occurred consistently with average weekly temperatures that ranged from 12.9 to 25.7 C. This was not unexpected because the house fly flight temperature threshold is approximately 13 C (Taylor 1963). Although the last three weeks in 2008 had w eekly average temperatures above the house fly activity threshold, it is possible that the accumulation of low temperatures and shorter daylight hours inhibited dispersal behavior. This is reflected in the very low recapture

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108 rates following week 3 and 5 re leases. Because week 5 was 21 d in duration, the last three collections of the 2008 study period represented a 5wk calendar period. A further challenge of dispersal studies is the diffusion (dilution) of marked insects across a landscape as the distance from the release site increases (Schoof and Siverly 1954b, Stein 1986). The capture of one marked fly 3 km from the release site could be equivalent to capture of several house flies at the release site, because the density of traps present at more distant concentric distances was less over a greater geographic area per trap than traps located more closely to the release site. Schurrer et al. (2004) determined that captures of house flies diminished with increased distance from the release site. House flies a re attracted to a wide variety of substrates, and every time a fly lands on an attractive substrate, the number of flies moving away from the release site is diminished (Murvosh and Thaggard 1966). Schoof and Siverly (1954b) estimated that approximately 50% of flies that land on an attractive site might subsequently progress towards an urban community that contains numerous attractants, so that the number of flies decreases by half at each visited site between the dairy and the town. The fact that such a small number of flies were recovered in town is probably a result of the manner in which the numbers of marked flies spread out spatially, thereby diluting the likelihood of recapture as they moved further from the release site (Quarterman et al. 1954, Sc hurrer et al. 2004, Krafsur et al. 2005). An estimated total of 37,200 marked flies were released during this study. The release of more marked flies, in theory, should increase the likelihood of capturing a marked fly on the far side of the town. Thimijan et al. (1972) increased their recapture rates from 0.17% (3 traps) to 0.51% (44 traps); however, their study was conducted in a closed barn so that an increased

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109 number of traps would likely influence recapture rates in their study more than placement of a dditional traps in my 3 km outdoor study area. If the marked flies in this study we re dispersing from the release site in all directions as reported in previous studies ( MacLeod and Donnelly 1960, Pickens et al. 1967), then the numbers of flies moving tow ards the town after a release represents only a portion of the released flies. I placed traps and recaptured marked flies in only one eighth (S S W portion) of a full 360 circle to study the dispersal of flies from the dairy into town. If the flies were dispersing equally in all directions (undirected) then 106 and 144 recaptured flies in 2008 and 2009 respectively, would be equivalent to a recapture rate of 848 (9.2%) and 1,152 (4.1%) in all eight ordinal directions. Dispersal of the house flies in multip le directions was very lik ely in the current study area, because there are several nearby dairies and poultry farms surrounding the release dairy. I f dispersal of flies was nondirectional then attractive sites in town that are sufficiently attractive to i nterrupt flight might receive a disproportionate number of flies from the dairies (MacLeod and Donnelly 1960. The effect of wind speed and direction upon house fly dispersal has been examined with varying results by previous researchers. Flies have been r eported to be blown long distances. Bishopp and Laake (1921) cite wind assisted dispersal of flies for distances ranging from 3.6 km (Hodge 1913) to a remarkable distance of 153 km (Ball 1917). During this study, monthly prevailing wind directions were southerly overall for the entire study. In 2008, prevailing winds were initially SSE and changed to SSW. In 2009, prevailing winds for June, July, August, and September were W, SW, SSW, and SE, respectively (Weather Underground 2009). Although in coastal area s of Florida, sea -

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110 breeze fronts can influence wind direction, central Florida is too far from coastal fronts to have any influence most of the time (Jones et al. 1991). The town was SW of the dairy release site, so for all months, fly dispersal was at least partially into the wind. In this situation, fly dispersal from the dairy into town would not passively rely on wind. Evidence that flies actively travel into the wind or at right angles to the wind using olfactory and optomotor senses for direct flight rather than being passively or actively transported by the wind was noted by Bishopp and Laake (1921) and Johnson (1966). Such movement would have resulted in fli es dispersing towards the town. The discrepancy between directed and undirected dispersal co ncepts suggests that a betterdesigned study on the potential of fly pathogen transmission into communities is needed to implement improved fly control programs at animal rearing facilities. The scope of such a study would need to include all house f ly pro ducing facilities within 6 km. However, each of those facilities also would share a fly population with additional sites located within another 6 km distance, making such a study impractical and perhaps impossible. Fly density could have been a dispersal factor in my study, because increased population density leads to increased displacement (Stein 1986), and by laboratory rearing of dairycollected progeny followed by release of those flies at the dairy, the density of the house fly population may have be en artificially increased. However, adult fly density on the dairy was not estimated, so the effect of population density upon recapture rates cannot be determined in the current study. When recaptured numbers of flies were regressed against the number of released flies, there was a linear relationship for both years combined, and for 2009. However, there was no relationship in 2008. This

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111 implies that the numbers of flies that were released, including the laboratory reared flies, might have influenced the overall dispersal rates or patterns of the representative fly population in 2009, but not in 2008. Capture of marked house flies 8 to 14 d after traps were placed demonstrates the ability of some marked flies to survive for longer than 7 d (Table 23), w hich confirms the findings of previous studies (Lindquist et al. 1951; Schoof and Siverly, 1954a,b; Pickens et al. 1967). Recapture of marked flies for up to 814 d after their release also provides information for how long any adult fly, whether marked or not, lives in the environment This is probably highly variable and is not known with certainty. Survival of marked flies for more than 7 d also implies that flies might disperse from breeding and aggregation sites on different days during their lifetime. While placement of traps for a 7 d period might be sufficient to capture most marked flies, it clearly is not long enough in all cases, as adult marked flies have been recaptured 10 d (Schoof and Siverly 1954a) and 20 d (Lindquist et al. 1951) after relea se, and pupal emergent marked flies have been recaptured 18 d after release (Pickens et al. 1967). In the current study, survival of marked house flies for more than 7 d was only recorded during weeks 4 and 7, so most house fly dispersal likely occurred wi thin 7 d after release. This is consistent with 7 d mean recovery rates of 84.7% and 95.5% in two field trials conducted by Pickens et al. (1967). Because most house flies seem to disperse within 7 d, it is likely that the principle potential health threat posed by house fly transmission of disease causing pathogens might be heavily concentrated within this time period. However, the ability for a few flies to survive for longer periods, combined with their continuous, random dispersal (Schoof 1959) from a release site to adjacent dairies

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112 and into urban areas could result in successful pathogen transmission over an extended period of time, so that control of house fly populations should include measures that are effective for longer than 7 d. In my study, f lies that had been in the environment for lo nger than 7 d were recaptured in wk 4 (7 November 2008) and wk 7 (21 May 2009). Therefore, survival in the environment past 7 d does not appear to be limited by seaso n (Table 2 1). Daily replacement of trap sleev es over a 2 wk period would provide more thorough information for daily dispersal rates and total recapture percentages. Increased trapping periods should be accompanied by incorporation of additional dust colors, so any flies that might survive 15 d or more could be observed. Flies should be caught, marked and released early enough in the day to promote fly activity, including dispersal. Pickens et al. (1967) observed that marked house flies tended to rest for 1530 min on buildings, fence posts and trees near the release site before dispersing. If releases are done too late in the day, then flies may not groom adequately or disperse due to decreased metabolism and avoidance of nighttime flight. Flies in this study were released at approximately 3:00 p.m. The weekly number of marked and released house flies ranged from 0 (no release due to low fly numbers or weather conditions) to 10,000. The total number of flies marked and released in this study (37,200) was low in comparison to those of previous mark r eleaserecapture studies: e.g., 171,427 (Schoof and Siverly 1954b) and 160,000 (Nazni et al. 2005). However, my weekly release levels were similar to Quarterman et al. (1954) who marked and released 13,500 flies, and Thimijan et al. (1972) who marked and r eleased 5 groups of 2,5005000 flies. However, the test area used by Thimijan et al. (1972) was a closed barn (17 x 10 x 3 m) that restricted fly dispersal. This study's 0.50%

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113 marked house fly recapture rates are higher than the filth fly recapture rates o f 0.0140.029% (mean 0.022%) obtained by Tsuda et al. (2009). When examining the Yates et al. (1952) data following exclusion of their highest capture trap which was placed adjacent to the release site, it is interesting to note that I obtained higher per trap recapture rates with pl acement of traps in only one or dinal direction (SW) away from the release site. Yates et al. (1952) placed their traps in eight ordinal directions. One can speculate from the relatively high mean recapture rates per trap during my week of highest release (wk 9) that house flies dispersed directionally towards the town. The presence of SSW prevailing winds during wk 9 further strengthens this hypothesis, because flies that dispersed towards town during wk 9 dispersed into the win d. This in turn implies that the role of house flies in potential pathogen transmission from dairies into urban communities might be more important than currently recognized Overall prevailing winds during the entire study were southerly, so that dispersa l of recaptured flies was probably not wind assisted, but attractant seeking or odor driven instead. Identification by dispersing flies of attractive sites in urban communities is likely a factor in house fly dispersal into these areas. Dispersal distances and recapture rates might be influenced by the type of flies used, i.e., fieldcollected or laboratory reared, as mentioned briefly above. Previous studies have indicated that use of field collected flies is more representative of dispersal under natural conditions than flies that are reared for multiple generations in the laboratory. Eddy et al. (1962) recaptured a 10fold higher percentage of fieldcollected flies than laboratory reared flies, implying that laboratory colonies may lose the ability to di sperse. Although laboratoryreared flies might lose the ability to disperse, their

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114 introduction in large numbers might increase the population of adult flies, causing increased dispersal of flies from the dairy, either of the natural population or of the newly introduced flies. It would be interesting to study the effect of large releases of flies upon existing house fly populations, to determine if natural dispersal rates match dispersal rates of recaptured marked flies. There were weather conditions that made collection of adult flies very easy, such as on days that storms were approaching when adult flies were very active in the air and remained hovering in the air in large numbers, even without human disturbance. This corresponds with previously repo rte d accounts that fly activity greatly increases under falling barometric pressure conditions associated with storm fronts (Wellington 1945, Holzapfel and Harrell 1968) Although there were no appreciable changes in overall barometric pressure (BP) during the entire study period, it is possible that changes in the barometric pressure in the hours preceding release of marked flies might have influenced house fly dispersal in this study. Barometric pressure decreases in an undul ating fashion and can only be observed with a column of mercury (Wellington 1945); thus, a net change in barometric pressure might be slight, yet still influence fly behavior. However, hourly weather data were not available, so that examination of the effect of weather for periods of less than 24 h was not possible. On 21 May 2009, the study area was experiencing its fifth consecutive day of steady, heavy rain that r esulted in extensive flooding. Adult f ly activity was negligible on this date. On 4 June 2009, fly behavior at both dairies consisted of hovering in large clouds approximately 23 m above the ground instead of the usual feeding and resting behavior. The daily average weather data obtained from the nearest weather station does

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115 not support these f ield observations. This is very likely due to the fact that the weather station was located approximately 29 km inland. Proper determination of weather influences upon house fly behavior should be tracked locally in future studies as distant weather stati ons do not provide data that always accurately responds to local microclimates. In Florida, adult house flies are primarily active mid day during spring and fall when maximum temperatures are cooler (Hogsette, personal communication). They adopt a diurnal activity behavior with increased summer temperatures, so that they become inactive during the hottest part of the day, and have two activity peaks: the first peak occurs in the morning and the second peak before dusk. During the 2009 portion of this study period, the daily temperatures steadily increased necessitating an earlier sweep net collection before the temperatures became hot enough to inhibit fly activity. Because the hot weather decreased adult house fly activity, another obstacle for sweep netti ng collection was created by flies congregating near the feed troughs that lined the edges of the barn. As the temperature became progressively hotter, the cattle spent increasingly longer periods in the shaded protection of the barn that also was equipped with large ceiling fans and mist sprayers that cooled the animals. When cattle were present in the barn, they had ad libitum access to the feed troughs, and therefore frequently had their heads in the grain where flies were located, making sweep netting d ifficult. Because cattle are easily frightened by sudden movements, in the interests of cattle safety, flies were not sweep netted at the feed troughs when cattle were present. The collection and rearing of immature flies and F1 rearing of adult flies gr eatly increased the number of house flies available for mark and release. In particular,

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116 laboratory rearing enabled the release of more uniform numbers of flies even when field populations were at relatively low levels. Release of mass numbers of laborator y produced F1 dairycollected adult flies could have changed dispersal behaviors for a couple of reasons. Increased larval density in the laboratory could produce smaller adults (Haupt and Busvine 1968), increase developmental time (Black and Krafsur 1986) or decrease adult emergence. These factors could delay adult emergence of earlier instars so that multiple age groups of flies entered the adult population simultaneously. Previous authors have noted that overcrowding led to decreased food consumption re sulting in smaller adult sizes (Haupt and Busvine 1968) with increased activity, and increased dispersal tendencies (Johnson 1969, Stein 1986). Smaller adult size may influence sexual maturity as well, so that small individuals might not be sexually mature Because sexual maturation of the ovaries inhibits or decreases the dispersal tendencies of the adult female fly (Johnson 1969, Stein 1986), it is possible that small stature could prolong the dispersal period past the typical 3 d previously reported for the house fly. Similar observations of reduced size and increased activity were previously described by Taylor and Sokal (1976), who also stated that larval overcrowding increases the tendency of adults to disperse. Initial efforts to maximize production of adult house flies from dairy substrates containing immature fly stages of all ages were very timeconsuming and inefficient, because of the emergence of adults in small numbers over a 1 3 wk time period. Allowing dairy collected adult house flies to ov iposit in the house fly diet increased the production of same age house flies that could be massreared using well established standard USDA house fly rearing procedures (Hogsette 1992)

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117 Provided that laboratory rearing did not alter oviposition behavior or larval mortality, the F1 generation produced by dairycollected adult flies should accurately represent the real fly population's relative mixture of sexes. However, age might not be accurately represented because natural populations have uneven distribution of different larval stages in the spilled grains at the dairy (Johnson 1966), whereas oviposition of dairy collected females generated a sameage population that was 13 d post eclosion. During the rearing process, larvae were observed clumping together in groups comprised largely of same age instars. This behavior could indicate diff erent nutritional or environmental needs or changing feeding capabilities of each life stage. However, this observed behavior in the laboratory might be changed from their behavior in the field, because they were reared under a different light:dark regime and were fed a different diet. One example of changing environmental needs for developing house fly larvae is seen when third instars migrate before pupating. While feeding larvae are active throughout relatively moist grain, aggregates of pupae are found in drier sections of the grain. Dairycollected house flies were observed to pupate just under the crust of spilled grains at the dairy. In the USDA laboratoryrearing facility, IR and F1 larvae mostly migrated completely out of the tray, and pupated in the folds of the cotton pillowcase used to cover the trays. Many larvae were also clumped together in the corners of the rearing trays. My dispersal study also may have been influenced by the age of released flies. Johnson (1966, 1969) observed that younge r flies disperse more readily than older flies. Because laboratory reared progeny of dairycollected flies were young flies aged 1 6 d, their dispersal activity might be higher than that of the typical dairy farm flies in the study area. In apparent contra st to Johnson (1966), Taylor and Sokal (1976) observed that

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118 older flies (3 6 d) dispersed readily while younger flies (13 d) did not disperse. However, Johnson (1966) noted that muscid flies disperse inter reproductively, i.e., after ovaries have matured, as well as prereproductively, and that females are more likely to disperse before egg development occurs, while gravid females are more likely to cease flight, and to land on suitable oviposition sites Similarly, Sasaki et al. (2000) observed that femal e flies aged 6 8 d were more dispersiv e than 6 8 d old males On a different note, Johnson (1966) states that populations, rather than dispersing, tend to mix within wide areas. Evidence of this is seen in the current study, because marked flies were recap tured at neighboring farms. On the one hand, the laboratory rearing methods used in this study may have produced well fed adults, as the nutrition level could have been greater in the GHFD media than in the dairys spilled grains. This might have resulted in "fatter" or healthier flies that may not have been inclined to disperse. Larval weights were not measured, so it cannot be determined which situation might have prevailed for either laboratory reared or dairy collected flies. Furthermore, it is possibl e that conditions changed from week to week in the laboratory reared flies, because the density of fly larvae in the GHFD was not measured. Eddy (1962) observed that laboratoryreared flies do not disperse as readily as field collected flies. However, Pick ens et al. (1967) did not observe any difference in dispersal rates or patterns between field collected and laboratoryreared flies. Conversely, larval crowding may have occurred in the trays, so that the house fly larvae were underfed and subsequently pr oduced undersized adult flies (Black and Krafsur 1986). Although I did not record fly weights or sizes for any fly stages used in this study, I did observe larval overcrowding during production of F1 fieldcollected

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119 progeny for release on 4 June 2009. Duri ng week 2, the F1 progeny exhausted the GHFD media, and half of each tray's larvae and exhausted media were transferred into clean trays containing fresh GHFD. Exhausted media containing flies was spread on top of fresh GHFD. The flies moved into the new m edia. The adult flies resulting from this production took longer to develop (19 d) than the USDA colony flies (14 d), appeared smaller, and were noticeably more active than sameage laboratory colony flies (personal observation) which were maintained in a n adjacent USDA rearing cage. Therefore, the laboratory rearing methods used in this study probably served, in at least one release week, to increase the dispersal behavior of the IR and F1 flies. This could have occurred if population pressure was placed on the house fly populations upon release due to mass introduction of thousands of flies. It might also have occurred because the individual flies were more inclined to disperse due to effects of larval overcrowding (Taylor and Sokal 1976, Black and Krafsur 1986). Increased dispersal rates due to these factors may have contributed to the high 0.81% (81/10,000) recapture rate of flies that had been released on 4 June 2009, when 40% of the released flies were F1 laboratoryreared progeny. C onversely, transpor t of laboratory reared progeny to the field in cages containing mixed sexes of flies aged 1 6 d with food and water provided ad libitum may have decreased dispersal tendencies, as many flies may have mated and obtained protein meals needed for egg developm ent. Environmental conditions such as temperature, precipitation, barometric pressure, wind speed and wind direction were evaluated individually to determine their impact on the dispersal of marked house flies. There may be a relationship between the numb er of flies that were marked and released and the recapture rate, although these data were not

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120 statistically analyzed in this study. There does not appear to be a relationship between precipitation and recapture rates. These results agree with those of previous researchers who also did not find a correlation between weather and house fly behavior (Eddy 1962, Lysyk 1993). Under natural conditions, there are many age groups of adult house flies at breeding and aggregation sites. It is unlikely that all adult flies disperse from these sites, and dispersal may be dependent upon house fly sex and age. The female to male ratio of adult flies collected in sweep nets may have influenced the dispersal rates and patterns, and female and male adults may rest in differ ent preferential sites during the day (Avancini and Silveira 2000); therefore, sweep netting in midday over feed troughs may have resulted in predominant collection of females. Events occurred on the dairy that influenced fly populations sizes, larval de velopment, and behavior. Mechanical fly control efforts included sanitation and habitat elimination, such as the removal of spilled grains, drainage of manure lagoons, and application of manure slurry to cropfields. Chemical control efforts included frequ ent application of permethrin to the backs of cattle, intermittent application of imidacloprid containing fly baits and intermittent application of permethrin around the exterior of the barns and milking parlors. The dairy also utilized pyrethroidimpregna ted ear tags on the animals. However, ear tags are unlikely to have decreased house fly abundance on the dairy due to fly biology and behavior. Several interesting results were observed in this study: 1) house flies dispersed 3 km from a dairy to a restau rant in a nearby town; 2) recapture rates (1.15% in 2008 and 0.50% in 2009) were comparatively high in this study; 3) recapture occurred at sites

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121 located within patchy habitats, implying direct flight over obstacles or windassisted dispersal; 4) recapture occurred at sites along major roads that provide corridor movement, implying possible landmark orientation or vehicle assisted transport; 5) most marked flies were recaptured within 7 d; and, 6) recapture of one fly (0.2%) in week 4 and seven flies (0.12% ) in week 7, despite zero releases the previous week, document fly survival and dust retention for between 8 and 14 d. The numbers of house flies recaptured in this study are probably under reported because house flies disperse in all directions (Pickens et al. 1967). The potential for house fly transmission of pathogens from dairies into town is clearly demonstrated by recapture of at least one fly at a restaurant in town. Because dairy cattle are the primary reservoir for enteric pathogens such as E. col i O157:H7 (Dunn et al. 2004b, Nmorsi et al. 2007), and this pathogen can remain viable on house fly exteriors (De Jesus et al. 2004) and in house fly guts for up to 4 d (Kobayashi et al. 1999, Sasaki et al. 2000), the potential for disease occurrence is tr emendously increased when house flies successfully disperse into communities from dairies. In that dairies provide both the source and dispersal mechanism for pathogen transmission, this study shows the importance of pathogen and house f ly management that dairy operators should consider. This also documents the need for additional efforts to educate and encourage producer utilization of available control methods to decrease house fly populations.

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122 Figure 21. Alsynite trap (Olson Products Inc., Medina, O H) placed at dairies and used to recapture on dairy and dispersing house flies.

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123 1.0 km 3.0 km 1.5 km 0.5 km 2.5 km 2.0 km 3.5 km Figure 22. Alsynite trap (Olson Products Inc., Medina, OH) locations and distance (km) from release point. Trap 1 is located at the release point. Traps that are located w ithin less than 0.25 km of a concentric circle are considered to be located at that radial distance. Trap distances are as follows: Traps 3, 4 and 5: 0.5 km; 6 and 20: 1.0 km; 7, 8, 24, and 25: 1.5 km; 9 and 21: 1.75 km; 10, 18 and 22: 2.0 km; 11, 17 and 23: 2.5 km; 13, 15 and 16: 3.0 km; 12 and 14: 3.5 km. Traps are placed at approximately 0.5 km intervals radiating out from the release site at the dairy, to the town which is located W SW of the dairy. Slight displacements necessary to accommodate roads, p rivate property, and other obstacles. Traps placed along major corridors such as roads, in edge habitats between open fields, and along shrub/tree lines.

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124 A B C Figure 23. Examples of dairycollected collected house flies following excessive tre atment with two dusts to determine 24 h mortality effects. A) flies dusted with corona magenta dust, B) flies dusted with arc yellow dust, and C) flies not dusted (control).

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125 Table 2 1. Mean and maximum distances flown per week and year by marked house flie s released at a dairy in north central Florida Distance (km) Week a Mean Max imum 1 0.01 0.10 2 0.29 1.00 3 0.35 0.50 4 b 0.00 0.00 5 c NA NA 2008 Total 0.22 1.00 6 0.39 1.50 7 0.73 2.50 8 1.12 1.75 9 0.73 3.00 10 0.10 0.10 11 0.64 1.50 2009 To tal 0.62 3.00 a Flies were collected during two study periods: wk 1 5 (23 October 2008 to 4 December 2008) and wk 611 (21 May 2009 to 25 June 2009). b Wk 4, marked flies recaptured only at release site. c Wk 5, no marked flies recaptured (NA, not appli cable).

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126 Table 2 2. Trap distances (km) from the release site the total numbers of marked house flies captured per alsynite sticky trap, and the cumulative percentage of marked flies captured across all study weeks from the dairy release site to the more distant traps placed in a rural n orth central Florida landscapea. 2008 2009 Test (wk) Test (wk) Distance from Release Site (km) Trap No. b 1 2 3 4 5 Marked Flies Cum. % Flies cb 6 7 8 9 10 11 Marked Flies Cum. % Flies b 0 1 17 1 0 1 0 19 17.9 11 1 0 52 6 1 71 51.1 0.1 2 27 56 2 0 0 85 98.1 0 4 1 11 3 2 21 63.8 0.5 3 0 0 0 0 0 0 25 0 0 9 0 0 34 88.9 4 0 0 1 0 0 1 99.1 0 0 0 1 0 0 1 5 0 0 0 0 0 0 0 0 0 0 0 1 1 1.0 6 0 0 0 0 0 0 0 0 0 0 0 1 1 92.8 7 0 0 0 0 0 0 0 0 0 0 0 0 0 20 0 1 0 0 0 1 100.0 0 0 0 0 0 0 d 0 1.5 8 0 0 0 0 0 0 0 0 0 1 0 0 1 93.8 24 NA d NA 0 0 0 0 1 0 0 1 0 1 3 25 NA NA NA NA NA NA 1 1 1 0 0 0 3 1.75 9 0 0 0 0 0 0 0 0 0 0 0 0 0 97.2 21 NA NA 0 0 0 0 0 0 1 4 0 0 5 2.0 10 0 0 0 0 0 0 0 0 0 1 0 0 1 97.9 18 0 0 0 0 0 0 NA NA NA NA NA NA NA 19 0 0 NA NA NA 0 NA NA NA NA NA NA NA 22 NA NA 0 0 0 0 0 0 0f 0 0 0 0 2.5 11 0 0 0 0 0 0 0 1 0 0 0 0 1 100.0 17 0 0 NA NA NA 0 NA N A NA NA NA NA NA 23 NA NA 0 0 0 0 0 0 0 0 0 0 0 3.0 13 0 0 NA NA NA NA NA NA NA NA NA NA NA 15 0 0 0 0 0 0 0 0 0 f 1 0 0 1

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127 2008 2009 Test (wk) Test (wk) Distance from Release Site (km) Trap No. b 1 2 3 4 5 Marked Flies Cum. % Flies cb 6 7 8 9 10 11 Marked Flies Cum. % Flies b 16 0 0 0 0 0 0 NA NA NA NA NA NA NA 3.5 12 0 0 0 0 0 0 0 0 0 0 0 0 0 100.0 14 0 0 NA NA NA NA NA NA NA NA NA NA NA Total 44 58 3 1 0 106 100.0 38 7 3 81 9 6 144 100.0 a Flies were collected during two periods: wk 1 5 (23 October 2008 to 4 December 2008) and wk 611 (21 May 2009 to 25 June 2009). b S ome traps were placed along main roads (3, 6, 7, 9, 10, 11, 22), while others were located in patchy habitats (4, 5, 8, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 23, 24, 25). Traps 1 and 2 were located outside opposing ends of release farms feed barn. c Cumulative percentage of marked flies captured from the release site to more distant traps. Traps (11 17, 23) located at were located within the town. Traps not in town were considered rural. d Trap 20 was missing. e NA, not applicable. Traps not used for collection were eliminated from any analysis. f Traps fell onto their sides on the ground.

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128 Table 2 3. Weekly recapture rate of marked house flies on alsynite sticky traps as a percentage of the number dusted and released the previous week at a dairy in north central Florida Test (wk)a Release Date No. Dusted & Released %SN:IR:F1 b Dusted Colorc Collection Date Duration (d) No. Marked Flies Collected Recaptu red Color % Recapture e 1 10/16/08 3,700 100:0:0 AY 10/23/08 7 44 AY 0.08 2 10/23/08 3,000 100:0:0 CM 10/30/08 7 58 CM 1.93 3 11/ 01/08 500 100:0:0 AY 11/07/08 6 3 AY 0.60 4 11/07/08 0 NA NA d 11/13/08 6 f 1 AY 0.20 5 11/13/08 2,000 100:0:0 CM 12/04/08 21 0 CM 0.00 2008 Total 9,200 100:0:0 NA NA 106 NA 1.15 6 05/14/09 6,000 100:0:0 AY 05/21/09 7 38 AY 0.63 7 05/21/09 0 NA NA 05/ 28/09 7 f 7 AY 0.12 8 05/28/09 2,000 50:50:0 CM 06/04/09 7 3 CM 0.15 9 06/04/09 10,000 60:0:40 AY 06/11/09 7 81 AY 0.81 10 06/11/09 6,000 50:0:50 CM 06/18/09 7 9 CM 0.15 11 06/18/09 4,000 100:0:0 AY 06/25/09 7 6 AY 0.15 2009 Total 28,000 71:4:25 NA NA 144 NA 0.51 NA, not applicable. a Flies were collected as follows : wk 1 5 (23 October 2008 to 4 December 2008) and wk 611 (21 May 2009 to 25 June 2009). b Estimated percentages of flies marked with fluorescent dust and released by each of three method s described in the text. SN, adult flies were sweep net captured, marked and released same day; IR, Immature flies were dairy captured and laboratory reared, then marked and released after adult emergence; F1, progeny were laboratory reared from ovipositing dairy collected adults, then marked and released after adult emergence. c Dust color: AY = arcyellow; CM = corona magenta; NA = no dust applied. d Percent recapture = (No. marked house flies recaptured divided by the no. house flies dusted and released ) 100. e No flies were marked and released in week 4 or in week 7. f Flies collected in weeks 4 and 7 were recaptured from previous releases in weeks 3 and 6 respectively. These data were not used in any analysis.

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129 Table 2 4. Total number of house flies and marked house flies captured on alsynite sticky traps following release at a dairy farm in north central Florida Capture data are normalized to number of flies per trap per day to adjust for different collection period lengths. Total House Flies Ca ptured a Marked House Flies Recaptured Wk Date Collected Duration b (d) Total Daily Mean c (flies/day) Trap Mean d (flies/trap/day) Total Daily Mean c (flies/day) Trap Mean d (flies/trap/day) 1 10/23/2008 7 4,655 665 33 44 6.3 0.3 2 10/30/2008 7 3,096 442 22 58 8.3 0.4 3 11/7/2008 6 2,847 475 79 3 0.5 0.0 4 11/13/2008 12 1,702 142 14 1 0.1 0.0 5 12/4/2008 21 841 40 2 0 0.0 0.0 2008 Total 13,141 353 30 106 3.0 0.2 6 5/21/2009 7 15,946 2,278 127 38 5.4 0.3 7 5/28/2009 14 12,632 902 100 7 0. 5 0.0 8 6/4/2009 7 3,101 443 25 3 0.4 0.0 9 6/11/2009 7 9,697 1,385 77 81 11.6 0.6 10 6/18/2009 7 2,867 410 23 9 1.3 0.1 11 6/25/2009 7 4,192 599 33 6 0.9 0.0 2009 Total 48,435 1,003 64 144 3.3 0.2 Grand Total 61,576 250 All weeks excep t weeks 3, 4 (6 d) and wk 5 (21 d) consisted of 7 d collection periods. Number of traps placed weekly: 2008, 20 traps; 2009, 18 traps. a Total house flies captured includes first time capture of unmarked house flies and recapture of marked and released ho use flies. b Flies collected in wk 4 and 7 were recaptured from previous releases in wk 3 and 6 respectively, indicating more than 7 d longevity in the field. These data were not used in any analysis. c Daily means (no. flies/d) for each test period were calculated by dividing total capture numbers by the duration (d) of that period. d Trap means (no. flies/trap/d) were calculated by dividing daily mean house fly capture numbers by the number of traps in use, i.e., wk 1 5, 20 traps; for wk 611, 18 traps.

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130 Table 2 5. H ouse fly release week and recapture rate and associated weekly weather data obtained from Weather Underground station approximately 30 km from the release site in north central Florida Temperature (C) Precipitation Wind Speed (km/h) Wind Di r Barometric Pressure (hPA) Wk Recapture Rate (%) Max Min Mean Total (cm) Mean Max Prevailing Min Max Mean 1 0.08 30.1 19.4 24.8 0.38 9.7 24 ENE 1006 1033 1019 2 1.93 26.3 12.6 19.5 0.04 9.7 24 ENE 1006 1035 1020 3 0.60 22.1 7.2 14.8 0.24 9.7 24 WSW 10 07 1033 1020 4 0.20 22.1 7.2 14.8 0.46 9.7 24 WSW 1013 1018 1020 5 0.00 25.0 9.5 17.5 0.76 8.1 24 SSE 1001 1031 1016 6 0.63 20.6 5.7 12.9 0.30 8.1 27 WSW 1013 1023 1018 7 0.12 25.2 17.0 21.3 0.46 12.9 35 ENE 1009 1023 1016 8 0.15 29.5 19.4 24.7 0.46 8 .1 27 SSE 1009 1023 1016 9 0.81 31.3 19.8 25.7 0.33 8.1 27 SSW 1011 1015 1016 10 0.15 30.6 20.5 25.6 0.57 6.4 29 WSW 1011 1016 1013 11 0.15 33.4 21.7 27.7 0.63 8.1 30 SSE 1005 1016 1013 W k 15 (23 October 2008 to 4 December 2008) and wk 611 (21 May 2009 to 25 June 2009).

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131 CHAPTER 3 ESCHERICHIA COLI O157:H7 PREVALENCE Introduction Escherichia coli O157:H7 is a bacterial pathogen that causes dysentery and diarrheagenic diseases in humans, but is commensally present in its primary reservoir, cattle. Es cherichia coli O157:H7 has been isolated from house flies, Musca domestica L., on dairies (Alam and Zurek 2004) Florida has a large number of dairies that are located in close proximity to human population centers, and the expansion of many urban areas ha s decreased the distance between dairies and towns. House flies typically disperse within 3.3 km (Parker 1916) to 5 km (Peppler 1944), although they move freely within urban areas for up to 6.7 km (Quarterman et al 1954). Additionally, house flies are capable of flying as far as 8 km from their breeding sites (Bishopp and Laake 1921). The ability of house flies to disperse within and between rural and urban areas indicates that house flies have tremendous potential to serve as vectors for transmitting E. co li O157:H7 from the dairies to the town centers, particularly to restaurants. The close proximity of dairies to towns might permit the pathogen to cause human disease by the fecal oral route through involvement of the house fly. Although E. coli O157:H7 ha s been isolated from dumpsters outside restaurants in Gainesville, FL (Butler et al. 2010), no study has been done in Florida to determine the potential of pathogen transmission from dai ries into towns by house flies. Diarrheal diseases impart an enormous toll on human and agricultural animal (e.g., cattle) populations, with severe health and economic impacts. Hospital expenses for human patients with infectious diarrhea can be four times greater than those of other patients. Similarly, medication expenses can be four times higher and the length of

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132 hospitalization can be three times longer (Suda et al. 2003). Economic impacts include lost incom e for family members who must miss work, as well as lost profits for employers (Buzby et al. 1996, Abe et al. 2002, Sapers and Doyle 2009). For cattle, increased operating expenses are incurred by dairy farms, feedlots and cattle rendering plants, sanitation efforts, including fly surveillance and management measures to comply with foodsafety federal regulatory manda tes (CDC 2009). Lawsuits can result in imposition of stiff fines upon many different foodrelated industries including producers, distributors, and restaurants. A recent example: in June, 2008, a lawsuit led to a $13.5 million settlement after a child in M ilwaukee, Wisconsin died due to consumption of E. coli O157:H7contaminated food (Powell 2008, Rohde, 2008). Enteric bacteria such as E. coli O157:H7 can be transmitted by the house fly, M. domestica and other flies (Greenberg 1971, Greenberg 1973) Cattle are a primary reservoir for E. coli O157:H7 (Heuvelink 2003), and dairy farms located near towns or residential areas may present a potential public health threat if pathogen persistence and high populations of filth flies co occur and if flies were to disperse from the dairy House flies typically disperse no further than 0.31.2 km (West 1951, Quarterman et al. 1954, 1964, Stein 1986, Mi lio et al. 1988, Alam and Zurek 2004), although they can travel as far as 8 km (13 mi) from their breeding sites (Bishopp and Laake 1921) I n a recent study at a dairy in n orth c entral Florida, I determined that house flies dispersed up to 3 km into a nearby town (Chapter 3). These characteristics of house fly behavior increase the potential for the introduction of pathogens from cattle to humans, particularly from sites conducive to flybreeding (e.g., dairies) (Kaufman et al. 2005b) to ne arby human population centers.

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133 Isolation and identification of E. coli O157:H7 by standard microbiological methods is time consuming and labor intensive. Because infection with E. coli O157:H7 is a nationally notifiable event in the United States (Mead an d Griffin 1998), detection methods that are rapid, selective and sensitive are important (Ogden et al. 2001). Szalanski et al. (2004) developed a 6 h protocol for detection of E. coli O157:H7 from house flies by polymerase chain reaction (PCR) instead of standard microbiological techniques. Many researchers use PCR to definitively confirm presence of virulence factor genes after presumptively confirming species identification by means of multiple microbiological and biochemical tests (Buma et al. 1999, Cagney et al. 2004, Alam and Zurek 2004). Usage of PCR techniques following microbiological methods improves identification and characterization of E. coli O157:H7 because more than 100 E. coli serotypes produce Shiga like toxins (Cebula et al. 1995). Presumptive identification of E. coli O157:H7 by microbiological methods can eliminate many serotypes prior to PCR analysis, and PCR can subsequently be used to confirm identification and to serotype additional characteristics of E. coli O15 7:H7. Use of PCR in place of further biochemical tests can save both time and money in the laboratory, while increasing isolation sensitivity. The pathogenicity of enterohemorrhagic E. coli O157:H7 is dependent upon possession of several virulence factors that are encoded in the genome or in a large plasmid. P CR provides a more selective and sensitive method to identify E. coli O157:H7 in samples (Visetsripong et al. 2007), and permits serotype differentiation (Beutin et al. 2007), and enables a faster ide ntification of E. coli O157:H7 than direct culture methods, which is especially important during outbreaks. Primer pairs can be used in uniplex or

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134 combined in multiplex PCR to detect and amplify target gene fragments of specific virulence factor genes, suc h as the rfbEH7 and fliCO157 genes. In this study the prevalence of E. coli O157:H7 in house fly, grain, and manure samples from two dairies and in house fly samples from two restaurant garbage dumpsters in a nearby town was determined by direct culture a nd confirmed by PCR detection of the rfbEH7 and fliCO157 genes. Materials and Methods The overall study area was the same as that described previously (Chapter 2). House fly populations were monitored at the feed barns of two dairies, A and B, with Dairy B located 1.5 km east of Dairy A. Samples were collected for microbiological analysis from four sites: both dairies and two restaurants, C and D, located 3 km and 3.5 km southeast of Dairy A, respectively. Collection sites were chosen following selection of a town in northcentral Florida that met all of the following criteria: 1) one or more dairies located within 3 km of the town; 2) one or more restaurants in town with a dumpster located adjacent to the restaurant; and, 3) a nearby residential human population. This research model represents a reasonable set of naturally existing conditions under which a potential for pathogen transmission from dairies to restaurants exists. These conditions allowed E. coli strains collected at dairies and restaurants to be compared. Three laboratories in Gainesville, Florida, were used during this study. The Food and Environmental Toxicology (FET) Laboratory at the University of Florida was used to isolate and identify E. coli O157:H7 from grain, manure, and pooled house fly samples using direct culture methods. The Veterinary Entomology (VE) Laboratory at the University of Florida was used to store untested samples for future analysis by

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135 polymerase chain reaction The United States Department of Agriculture, Agricultural Research Service, Center for Medical, Agricultural and Veterinary Entomology (USDA ARS CMAVE) laboratory was used to measure house fly population counts as described below. House fly population trends were monitored weekly during selected intervals at both dairies from 7 June 2008 to 23 September 2008 using sweep nets (i.e., small "butterfly" nets), Scudder grids (Scudder 1947), a portable slatted wood frame placed on top of fly breeding areas, and fly spot cards (Lysyk and Axtell 1985), 7.5 x 12.5 cm index cards placed on walls on which flies excrete and regurgitate while resting (see Chapter 2 for a full description). Use of multiple monitoring methods was performed with the intention of examining these data for possible trends between these three monitoring methods (Table 31) as well as for correlation between house fly populations and E. coli O157:H7 prevalence. Nets, gr ids and cards were all used as close to fly aggregation and breeding areas as possible, and collection locations were the same every w eek unless noted differently. "Snapshot active monitoring of house fly populations by sweep nets and Scudder grids was discontinued after 23 September 2008 with the cessation of microbiological sampling, but weekly spot card sampling was continued until 4 December 2008. Because spot card data may represent activity of multiple species of flies, sticky cards, index cards (7.5 cm x 12.5 cm) covered with adhesive on one side so that flies become captur ed when they land (see Chapter 2 for a full description), were placed along barn walls at Dairy A from 30 October 2008 to 4 December 2008. Sticky cards permitted differentiation between fly species. Thus the sticky card and spot card data were examined for potential

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136 trends or correlations of house fly populations on Dairy A, the release site in my dispersal study (Chapter 2). Alsynite traps were also placed at each dairy 1 2 m outside of the feed barns at each dairy from 16 September 2008 to 4 December 2008 as part of a hous e fly dispersal study (Chapter 2). Because some of the alsynite traps were in use at the feed barns when spot and sticky cards were in use, the alsynite traps provided an additional method of house fly population monitoring and allowed for examination of correlations between monitoring methods. House fly monitoring occurred again from 14 May 2009 to 25 September 2009 using spot cards, sticky cards and alsynite traps at Dairy A, and spot cards and alsynite traps at Dairy B. Sweep net monitoring of adult house f lies was performed at both dairies by sweeping an insect net (45 cm diameter, Mod. No. 7112NA, Bioquip, Rancho Dominguez, CA) in one figure eight pattern. Sweeps were performed so that the net was at a height of 0.5 m above adult aggregation areas in feed trough areas. Sweep nets were also performed above spilled grains that were below feed augers. Sweep nets were performed while walking towards the sun, to prevent disturbance of house flies due to casting a shadow over resting flies and at a fast walking pace. One to four net sweeps were performed at each dairy from 7 June 2008 to 23 September 2008. M ean sweep net counts for each dairy were calculated for each collection day by dividing the total number of flies captured by the number of sweeps performed. Small Scudder grids (45 x 45 cm) (S cudder 1947, Murvosh and Thaggard 1966) (Fig. 31) consisted of 12 slats of rough unfinished wood (8 mm x 45 cm) laid parallel to each other and spaced evenly along a 45 x 45 cm wood frame. This provided a series of alternating edges for flies to rest upo n. Scudder grids were placed on the ground on top of

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137 spilled grains under augers or on top of feed troughs 110 times at each dairy from 7 June 2008 to 23 September 2008. Each Scudder grid was photographed after it was in place for a period of 5 sec, and t he number of house flies on each grid was counted by examining digital photographs on a computer monitor at the USDA ARS CMAVE laboratory. Where photographs were not available, estimates were noted on the data sheet. Ten spot cards were placed at each dai ry inside a barn adjacent to the milking parlor (Dairy A, milk barn; Dairy B, south barn) on 5 August 2008. Because use of the milk barn at Dairy A discontinued in September 2008, spot cards were relocated to Dairy A's feed barn located approximately 0.25 km north of the milking parlor on 23 September 2008. Use of spot cards was continued in Dairy A's milk barn until 30 October 2008 to track fly population changes after departure of the cattle. Spot card monitoring at each dairy using Dairy A's feed barn and Dairy B's south barn continued until 4 December 2008, and was resumed from 14 May 2009 to 25 June 2009. Spot c ards were placed horizontally at a height of 23 m and were flush against either support beams or rafters. In all barns, spot cards were placed at an approximate height of 23 m and in locations where they were not within reach of the cattle Spot cards at Dairy B and at Dairy A's milk barn were inserted into metal frames nailed to the wooden support beams of these barns and held in place with one small binder clip placed at the top edge of the metal frames (Fig. 3 2) or by clipping them to large metal support beams with small and medium binder clips (Office Depot, Delray Beach, FL) (Fig. 33). Sticky cards (7.5 cm x 12.5 cm) (Hogsette et al. 1993) were used from 30 October 2008 to 4 December 2008, and again from 14 May 2009 to 25 June 2009. Fourteen sticky cards were placed at Dairy A's feed barn by clipping them with large paper clips to the

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138 steel support beams immediately below the spot cards so a gap of approximately 1 cm existed between the spot and sticky cards (Fig. 3 3). Three alsynite sticky sleeve traps (Broce 1988, Hogsette and Ruff 1990) were used for 5 wk from 30 October 2008 to 4 December 2008 in conjunction with my 2008 dispersal study (Chapter 2). Two traps were located at Dairy A and one was located at Dairy B ( Chapter 2, Fig. 21). All alsynite traps were placed within 1 2 m of a specific barn or feed trough as described below. The traps at Dairy A were placed 100 m apart, with one trap (trap 1) at the SE corner and the other at the SW corner of Dairy A's feed barn (trap 2). The trap at Dairy B was placed at the SE corner of the south barn (trap 8). Five traps were used at the dairies from 14 May 2009 to 25 June 2009 in conjunction with my 2009 dispersal study (Chapter 2), with two at Dairy A (traps 1 and 2) and three at Dairy B (traps 8, 24 and 25). Trap loca tions used in 2008 were used again in 2009. Two additional alsynite traps were placed on Dairy B during 2009: one (trap 24) was located W of the calf feed trough, approximately 0.3 km S of the adult barns, and the second (trap 25) was placed at the NE corn er of the adult barns approximately 100 m from the original alsynite trap location used in 2008. These alsynite traps were relevant to fly monitoring efforts because they were placed immediately outside of barns on both dairies where cattle are fed and where adult flies aggregate. In addition to house fly population monitoring, I collected samples to test for the presence of E. coli O157:H7 by direct culture microbiological analysis. Samples collected from dairies included adult house flies, spilled grains and fresh manure. At restaurants, only adult house flies from the garbage dumpsters were sampled.

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139 Flies were collected by sweep netting, with the additional use of sterilized materials. All materials that came into direct contact with flies were either purchased sterile, autoclaved for 60 min at 121 C, 15 psi, or placed in 6% hypochlorite (bleach) (Clorox Co., Oakland, CA) solution for 15 min prior to use. Sweep nets were autoclaved and remained sealed in autoclave bags until used to collect a single sample in the field. Each sample of flies collected by sweep netting was transferred to a sterile 120 ml clear polypropylene specimen cup (Model 70756, Samco Scientific Corp., San Fernando, CA). Fresh nitrile gloves (Best Glove, Inc., Menlo, GA), sterilized with 6% bleach (Clorox Co., Oakland, CA), were worn when using each sweep net. Specimen cups containing fly samples were placed on ice, in a large cooler (50 L, Igloo Corp., Houston, Texas) for approximately 1 min (1020 C), to chill flies without mortality. Up to 10 flies were gently shaken onto a chilled metal pan (lined with aluminum foil which was changed after every sample) and identified to species. Flies not removed from specimen cups for sorting were later identified in the laboratory. Up to 10 M. domestica were placed individually into sterilized snapcap microcentrifuge tubes (P/N 02 681 240, Fisher Scientific Co., Waltham, MA). Flies were manipulated using sterilized feather weight forceps (P/N 4750, Bioquip, Rancho Dominguez, CA). The 10 indivi dually contained flies were placed inside a prelabeled sealable, clear, plastic bag, and placed in the cooler. All fly samples remained in the cooler at 10 20 C for 26 h until removed for laboratory processing later that same day. During the cooler months of November April when fly populations were low, isolation of 10 individual flies was given priority over pooled samples; i.e., if only two house flies were collected, then each of the two was placed individually in sterile 1.5 ml

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140 cap microcentrifuge tubes. If 12 flies were collected, then ten were individually placed in microcentrifuge tubes and the remaining two flies were retained in the specimen cup. During the warmer months of MayOctober, flies collected at each site were transferred to sterile s pecimen cups, placed on ice in a cooler, transported to the laboratory, and knocked down by placement in 20 C for approximately 1 min. Afterwards, specimen cups were removed from the freezer, and 10 individual flies were transferred into individual micro centrifuge tubes as described previously. Cattle manure was obtained using sterile materials. Manure samples consisted of fresh droppings, and were obtained within 30 sec after animal defecation. Slurry samples were collected from barn floors or lagoons ( wastewater retention ponds). Grain samples were obtained from feed troughs or from spilled grains below feed augers. Manure, a nd grain (substrate) samples were obtained manually by two methods (Fig. 34). In the first sampling method, approximately 100 g of each substrate was scooped out with a sterilized metal spoon. Manure samples were obtained from the center surface of the fresh dropping. Manure samples were placed in prelabeled 120ml sterile specimen cups and placed inside the cooler (1020 C) for transport to the laboratory where they were processed within 24 h of collection. In the second sampling method, two sterile swabs, premoistened with buffered peptone water (BPW) (Oxoid Ltd., Basingstoke, Hampshire, England) or trypticase soy broth (TSB)(D ifco, Becton, Dickinson and Co., Sparks, MD), were inserted 2.55.0 cm deep into manure or grain, and rolled around briefly until completely coated with substrate particulates. Both swabs from each substrate were placed together as one sample in sterile 15 ml polypropylene centrifuge tubes (Model No. 352096, Becton Dickinson Labware, Franklin Lakes, NJ)

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141 containing 9 ml of sterile BPW or TSB. Swab samples were placed in a shaded location in the vehicle, and transported to the laboratory at ambient temperatu re (25 30 C). Air temperature, relative humidity (RH), and barometric pressure (BP) data were recorded onsite at Dairy A using a portable weather station (Model No. 00589W, Acu Rite, Jamestown, NY) and examined to determine if any of these weather condi tions influenced house fly population activity determined by house fly monitoring methods. At the time of sample collection, both the substrate and surface temperatures of manure and grain samples were measured. Substrate temperatures were measured by ins erting a digital soil probe thermometer (Model No. 6310, Spectrum Technologies, Inc., Plainfield, IL) 2.5 5.0 cm into the grain or manure. The digital thermometer was allowed to calibrate while samples were obtained. Surface temperatures were measured by pointing an infrared thermometer (Raynger ST2, Raytek Corp., Santa Cruz, CA) at the center of the sample from a distance of 1 m. All samples that were scheduled to be processed while fresh using standard microbiological methods were delivered to a Biosafet y Level 2 (BSL2) laboratory in the FET Laboratory within 6 h of collection, and processed within 24 h of collection. Fly samples were placed in a 20 C freezer for up to 30 min to reduce the potential for flies escaping during processing; substrate sample s remained in the cooler at 10 20 C, and swab samples were maintained at room temperature (25 C) until processed. Some samples were reserved by storage in a 20 C freezer at the VE Laboratory for future identification and serotyping by polymerase chain reaction (PCR). All samples were processed using standard BSL2 laboratory aseptic techniques, and in compliance with the University of Florida's (UF) Environmental Health and Safety

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142 (EHS) Biosafety Protocols (UF EHS 2008). Pipette tips, tubes, broths, and other reagents were either purchased sterile or were autoclaved for 60 min at 121 C, 15 psi, except where noted differently, using a table top autoclave (Sterilmatic STME, Market Forge, Ramsey, MN). Agar plates were poured and allowed to solidify inside a BSL2 cabinet. Samples were processed at an open BSL2 bench as described below. Aerobic plate counts (APCs) of unenriched background microbial (BM) organisms were performed. Each sample was vortexed for 3060 sec to resuspend bacteria and held at room te mperature for 1 min to permit debris to settle. A 1 ml aliquot of each unenriched sample was pipetted into a 15 ml (O.D. x Length 16x125 mm) borosilicate glass screw capped culture tube (No. 982516X, Corning Pyrex Inc., Lowell, MA) containing 9 ml of BPW. Serial dilutions were prepared for each sample by sequential pipetting 1 ml of the bacteria:broth mixture to tubes containing 9 ml BPW, for a total of six serial dilutions, 101 106. Two aerobic plate counts using Petrifilm APC plates (3M, St. Paul, M N) were performed for each dilution tube. First, tubes were vortexed for 3060 sec and allowed to settle for 5 10 sec. Then 1ml aliquots were pipetted onto APC plates. In this way, the resultant culture plate dilutions ranged from 101 106 colony formi ng units per gram (CFU/g). A new pipette tip was used for each tube, and dilutions were transferred sequentially from the most to the least dilute, to decrease the possibility of crosscontamination. After the 1ml aliquot of each dilution was placed on a n APC plate, a proprietary plastic tool (spreader) that was included with the Petrifilm APC plates was gently placed upon the APC plate to spread the diluted sample out until it covered a 20-

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143 cm2 surface. Plates were incubated at 37 C for 6 18 h and plate counts were performed the following day by counting CFUs on each plate. Plates with 14300 colonies were used to calculate average values for total CFU/g of aerobic bacteria for each sample. To improve selectivity for E. coli O157:H7 from complex fecal a nd decomposing organic matter containing high concentrations of competing background microorganisms, TSB was modified by the addition of novobiocin (20 mg/l) (mTSB+N) (FDACFSAN 2007a) for the enrichment broth (Desmarchelier et al. 1998). Bacteria species used as positive and negative controls in this study were obtained from Dr. Huang (Auburn University, AL) and are maintained by Dr. Simonne in the FET Laboratory. Nalidixic acid resistant E. coli O157:H7 strain 204P was used as a positive control. Negativ e controls used in this study were Shigella dysenteriae (ATCC 49550) and Salmonella thompsoni (ATCC 8391). Frozen fly samples containing multiple fly species were removed from the freezer, and up to 25 M. domestica were randomly selected and placed as a p ooled sample in mTSB+N. Species other than M. domestica were discarded. Adult house flies were incubated individually or in pools of up to 25 flies. Some thawing occurred during sorting. For pooled samples with up to nine flies, M domestica adults were ad ded to 9 ml mTSB+N in a sterile 15 ml polypropylene centrifuge tube (Model No. 352096, Becton Dickinson Labware, Franklin Lakes, NJ). Pools containing 10 or more adult flies were placed in a 50 ml polypropylene centrifuge tube (Model No. 43089, Corning In corporated, Corning, NY) containing 25 ml mTSB+N. Individual samples consisted of one M. domestica adult that was placed into a sterile 15ml polypropylene centrifuge tube

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144 (Model No. 352096, Becton Dickinson Labware, Franklin Lakes, NJ) containing 9 ml mT SB+N. Samples were incubated at 37 C for 24 2 h. Substrate samples were initially enriched using 25 g of each substrate. However, following challenges with odor in the FET Laboratory, a swab sample technique was adopted (Rice et al. 2003, Greenquist et a l. 2005, Davis 2006). In the initial substrate enrichment process, 25 g of each substrate sample were added to individual double bagged 400ml stomacher bags (177 mm x 305 mm) (Stomacher 400 Classic, P/N BA6041/CLR, Seward Co., Seward, UK) containing 225 ml of mTSB+N. The substrate mTSB+N mixture was homogenized in a stomacher for 60 sec. Unused portions of substrate samples from each bag were stored in a freezer at 20 C for future analysis. Some of the unused substrate samples were disposed of following mechanical failure of the freezer that resulted in thawing of samples. In the second substrate enrichment process, swab sample tubes (Model No. 352096, Becton Dickinson Labware, Franklin Lakes, NJ) containing 2 substrate inoculated swabs in 9 ml BPW were processed by vortexing each tube for 3060 sec. One swab and 1 ml supernatant was transferred to a new 15ml centrifuge tube (Model No. 352096, Becton Dickinson Labware, Franklin Lakes, NJ) containing 9 ml of mTSB+N. Transferred samples were incubated at 37 C for 24 2 h. The second swab and remaining BPW supernatant (8 ml) were stored unaltered in a freezer at 20 C for future analysis. Following selective enrichment, enriched samples were removed from the incubator, vortexed for 3060 sec and allowed t o settle for 1 min in preparation for enumeration and isolation procedures. Enumeration of E. coli O157:H7 was performed using SMAC plates supplemented with cefixime (15 g/l) and potassium tellurite (1.25 g/l) (CT SMAC)

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145 (Alam and Zurek 2004, FDA CFSAN 2 007b). One ml aliquots of each unenriched vortexed sample were transferred to screw capped culture tubes containing 9 ml of sterile BPW and vortexed for 3060 sec. Six serial dilutions were prepared for each sample, as previously described, and 100 l aliq uots were pipetted onto each of two CT SMAC plates, i.e., double plated, so that two plates were prepared for plate dilutions that ranged from 102 to 107. The unenriched bacteriabroth mixture was spread evenly over the plate using a glass rod. Spread pl ate counts of E. coli O157:H7 were obtained by averaging the number of CFUs for plates that contained 14300 colonies. CT SMAC spread plates were incubated at 37 C for 242 h. Isolation of E. coli O157:H7 was performed by immunomagnetic separation (IMS) and direct culture, as described below. Separation, following Invitrogen's proprietary protocol (Dynal 2007), was performed using magnetic E. coli O157specific antibody coated beads (Dynabeads anti E. coli O157, Invitrogen, Carlsbad, CA) that increased 8 beads/ml]) were aliquotted into a 1.5 ml microcentrifuge tubes and 1 ml of enriched house fly sample supernatant was added to each tube. Tubes were placed in a manually operated magnetic stand, (Ambion 6 Tube Magnetic Stand, Invitrogen, Carlsbad, CA), closed and inverted gently for 10 min at room temperature (RT) to permit the E. coli O157:H7 to bind antigenically to the beads while keeping the beads suspended in the supernatant. A magnetic plate was i nserted into the magnetic stand, and tubes were inverted multiple times to form a pellet of concentrated beads at the bottom of the tube, along the magnetized side of the tubes. Tubes were allowed to stand for an additional 3 min at RT to maximize recovery of beads coated with E. coli O157. Supernatant was pipetted out

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146 and discarded, taking care not to dislodge the bacteriabead pellet complex. The magnetic plate was removed and three sequential 1 ml 20X PBS Tween 20 (Thermo Scientific, Rockford, IL) wash buffer rinses were performed to resuspend the beads and enhance removal of nonspecific binding microorganisms. After the third rinse, the bacteria bead complex was plated onto a CHROMAgar (BBL CHROMAgar O157, Beckton Dickinson Co., Franklin Lakes, NJ) plate by mo istening a sterile swab with the bacteria bead complex and streak plating half the agar plate with the moistened swab. The second half of the plate was loopstreaked to enhance isolation of colonies. Plates were incubated at 37 C for 242 h. After incuba tion, both CT SMAC spread plates and CHROMAgar isolation plates were examined for presumptive E. coli O157:H7 colony growth. Presumptive E. coli O157:H7 appeared colorless with or without a light smoky center (sorbitol negative) on CT SMAC plates, and light violet to violet on CHROMAgar plates. Up to five presumptive positive E. coli O157:H7 colonies from CHROMAgar plates were selected and streaked onto CT SMAC plates which were incubated at 37 C for 24 2 h. Presumed E. coli O157:H7 colonies were subsequently loop streaked onto nonselective, general growth Trypticase soy agar with yeast extract (TSAYE) plates and incubated at 37C for 24 2 h. Sub cultures were prepared for longterm storage by transferring a loopful of one TSAYE colony into 850 l of a st erile Luria Bertani (LB) broth. The bacteria:broth mixture was pipetted into 150 sterile l glycerol in 2 ml cryogenic vials (P/N 1050026, Fisher Scientific, Pittsburgh, PA). The LB broth, bacteria, and glycerol were gently

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147 pipetted in and out until thor oughly mixed (510 times) without introduction of air bubbles. Finally, the bacterium and its 15:85 glycerol:bacteria broth mixture were vortexed for 30 s and stored at 20 C, to be available for further DNA analysis by polymer ase chain reaction (Chapter 5). Where sufficient samples were available, additional biochemical tests were conducted. These tests included phenotypic expression of characters typical for E. coli O157:H7: 1) hydrolization of tryptophan; 2) the presence of a brilliant green sheen on L EMB agar; and, 3) a lack of fluorescing 4methylumbelliferyl D glucuronide (MUG). Hydrolization of tryptophan was assessed by placing a filter paper wetted with Kovac's Reagent (Ricca Chemical Co., Arlington, TX) on each positive growth plate. Colonies that are presumptive positive for E. coli O157:H7 typically turn pink. Presumptive E. coli O157:H7 colonies were plated concurrently onto two different media; Levine's eosin methylene blue (L EMB) (Oxoid) agar, and fresh TSAYE agar plates. Incubation of L EMB plates, which inhibit gram positive growth, occurred at 37 C for 242 h. Following incubation, plates were examined for a distinctive bright metallic green sheen on dark blue to black nucleated colonies. This brilliant green sheen is characteristic o f E. coli O157:H7, and differentiates E. coli O157:H7 from nonpathogenic E. coli that turn dark green without the brilliant green sheen on L EMB agar. Isolates that were transferred to TSAYE plates were tested for the presence of gl ucoronidase an enzyme that hydrolyzes 4methylumbelliferylD glucuronide (MUG ) t o yield fluorescing 4 methylumbelliferone by addition of one ColiComplete (Bothell, WA) disk to the plate's heaviest bacterial streak. After incubation at 37 C for 242 h,

PAGE 148

148 colonies presumptive for E. coli O157 :H7 were considered negative if they lack ed blue fluorescence around the disks when examined under 365 nm ultraviolet (UV) light. Enumeration of aerobic bacteria was performed using Petrifilm APC plates containing 14300 CFU/g. Aerobic bacteria counts wer e obtained for 19 selected samples that consisted of nine house fly, seven grain and three manure samples that were collected from 31 May 2008 to 26 August 2008. Aerobic plate counts of the two grain and two house fly samples obtained from Dairy A on 26 August 2008 were averaged for each type. Enumeration data for the remaining 17 samples were based on single samples. When multiple dilution plates containing 14300 CFU/g were obtained from individual samples, the range of those plates was recorded. Presumpt ive E. coli O157:H7 colonies were counted using CT SMAC spread plates containing 14300 CFUs on two dates: 31 May 2010 and 14 June 2010. Samples of 31 May 2010 were enumerated, but not processed further. Samples of 14 June 2010 were submitted for microbiol ogical analysis after enumeration of E. coli O157:H7 was completed. Colonies that tested positive on CHROMAgar plates w ere classified as presumptive positive. However, due to failure of the CHROMAgar on 26 August 2008 and 16 September 2008, isolates that were presumptive positive on CT SMAC agar plates on 26 August 2008 and 16 September 2008 were also submitted to PCR. Thus, isolates of 24 samples of the 57 dairy collected samples were submitted to PCR with isolates from 11 samples originating from CHROMA gar plates and isolates from 13 samples originating from CT SMAC plates.

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149 M ultiple isolates were obtained from several samples due to sequential transfer of indi vidual colonies dur ing the microbiological testing. Therefore, prevalence rates were determined for both the number of samples tested and for the number of isolates that were sub cultured from samples. Sub cultures of each isolate were prepared for long term storage by aseptic loop transfer of one loop of logarithmic growth phase E. coli O157:H7 fr om TSAYE culture plates to 850 l sterile LB (Oxoid) broth (lysogeny broth; Bertani (1951, 2004)). The LB broth and bacteria were mixed by gentle pipetting and then added to 150 l sterile glycerol in a sterile 2 ml cryogenic vial (P/N 10500 26, Fisher Sc ientific, Pittsburgh, PA). The LB broth, bacteria, and glycerol mixtures were stored as a 15% glycerol stock in 20 C nonthawing freezer. Presumptive positive isolates were prep ared for DNA extraction and PCR analysis from 1 December 2009 to 16 April 2010, approximately 1.5 yr after placement in the freezer To prepare frozen isolates for DNA extraction and PCR, fresh cultures of each isolate ml culture tube containing 2 ml ste rile LB bro th. Transfer of bacterial stock was performed in a BSL2 c abinet. The stock was returned to the freezer to minimize thawing. Culture tubes containing fresh bacteria broth mixtures were incubated at 37 C for 242 h with shaking (250 RPM) (LabLin e Orbit EnvironShaker, Lab Line Instruments Inc., Melrose Park, IL). Eight hundred fifty microliters of this fresh culture were used to make another glycerol stock to be placed in a 70 C freezer for long term storage, and the remaining culture was used for DNA extraction and PCR amplification.

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150 Total genomic DNA was extracted from stored bacterial isolates originally obtained from house fly, grain and manure samples using the QIAquick DNeasy Blood and Tissue Kit (QIAgen, Valencia, CA). Extracted DNA wa Buffer AE and stored at 4 C. Polymerase chain reaction was performed using one multiplex and two uniplex PCR assays ( Table 3 2) to amplify gene fragments using two primer pairs (20 pmol/ml each) designed to target the rfbEH7 and f liCO157 genes (Hu et al. 1999, Cagney et al. 2004, Szalanski et al. 2004). The fliCO157 primer pair amplifies a 625 bp E. coli O157 serotype gene fragment (Gannon et al. 1997) and the rfbEH7 primer pair amplifies a 259 bp E. coli H7 gene fragment (Paton an d Paton 1998). Following low amplification of both gene fragments simultaneously in the multiplex PCR, samples were submitted to uniplex PCR for each gene fragment separately. The following three assays were conducted to test for the genes of interest: As say 1, Multiplex PCR for rfbEO157 and fliCH7; Assay 2, Uniplex PCR for rfbEO157; and Assay 3, Uniplex PCR for fliCH7. Master mix reagents for each assay consisted of deionized sterile water, 10X DNA Polymerase PCR Buffer ( MgCl2), dNTP Mix (10 mM each), 50 mM MgCl2 oligonucleotide primers (20 pmol/l each) (Eurofins MWG Operon, Huntsville, AL) (Table 3 2) and recombinant DNA polymerase Taq enz yme (Invitrogen, Carlsbad, CA). For the multiplex assay and for the amplification of the rfbEH7, the PCR progr am consisted of an initial denaturing at 94 C for 2 min, followed by 35 cycles of denaturing at 94 C for 45 sec, annealing at 56 C for 45 sec, and extending at 72 C for 1 min. A final extension was performed at 72 C for 5 min. The PCR program for ampl ifying

PAGE 151

151 flicCO157 gene region was similar to those just described but the annealing temperature was increased to 65 C. Sterile distilled water was used as a negative control in each assay. Bacterial positive and negative controls used in this study were o btained from Dr. Amarat Simonne at the University of Florida, in the Food and Environmental Toxicology Laboratory. Nalidixic acid resistant E. coli O157:H7 strain 204P was used as a positive control. Negative controls were Shigella dysenteriae (American Ty pe Culture Collection (ATCC) 49550) and Salmonella thompsoni (ATCC 8391). Cultures used f or this study were maintained in the laboratory of Dr. James E. Maruniak at the University of Florida, Entomology and Nematology Department, with subcultures stored at both 20 C and at 70 C. Amplified PCR gene fragments were visualized by ethidium bromide staining of the PCR product that was separated in 1% agarose gel electrophoresis. Visualized PCR products were photographed using ultraviolet (UV) light. Isolates were considered positive if both the fliC and rfbE gene fragments were amplified Because multiple isolates corresponded to each dairy collected sample, samples were considered positive if at least one isolate was positive. The percentage of positive samp les was calculated by dividing the number of positive samples by the number of samples that were submitted to PCR and multiplying by 100. DNA Quantification DNA concentration of fresh ly cultured isolates was obtained by measuring the absorbance of light a t wavelengths of 260280 nm, using a spectrophotometer (Nanodrop 1000 mini spectrophotometer, Thermo Scientific, Waltham, MA) or was estimated by visual comparison of the band intensity with the standard 100 bp ladder purchased from

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152 Invitrogen (Genvault 20 10). Spectrophotometry nanodrop analysis was performed in accordance with the manufacturers instructions in Dr. Michael Scharf's laboratory, Entomology and Nematology Department University of Florida. 16S rDNA PCR Analysis To ensure that PCR product ge ne fragments were from bacterial DNA, a broad range 16S rDNA PCR assay was performed on 46 selected isolates. Broad range 16S rDNA PCR analysis was performed in Dr. Volker Mai's laboratory, Emerging Pathogens In stitute, University of Florida. Statistical A nalysis Seasonal trends of house fly populations were estimated for the sampling period using weekly mean counts for both passive and active fly monitoring methods at both dairies during the respective weeks of placement of each passive monitoring method. Passive fly monitoring estimates were obtained wit h spot cards, sticky cards and sticky sleeves on alsynite traps. Active mon itoring estimates were obtained with weekly active "snapshot" counts of Scudder grids and sweep nets at four sampling sites. Hous e fly population data were subjected to PROC UNIVARIATE to examine normality and PROC MEANS to calculate means using SAS Version 9.1 (SAS 2002). Differences between sites or between treatments (monitoring methods) were determined by one way analysis of var iance (ANOVA) using Fisher's categorical test where populations were not normally distributed (PROC ANOVA) (SAS 2002). Correlations a mong sticky, spot and alsynite monitoring methods were analyzed using a three way correlation analysis (Pearsons coefficient) analysis with Dairy A data from wk 1523 when all three monitoring methods were in use (PROC CORR, SAS 2002). Paired correlations between spot and sticky cards were performed using Pearsons coefficient

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153 with Dairy B data from weeks 1523 when both methods were in use (PROC CORR, SAS 2002). Correlation analysis was conducted for spot card data collected during weeks that cards were in use at both dairies. Scudder grid and sweep net counts were subjected to correlation analysis (PROC CORR, SAS 2002). Mete orological data (BP, RH and air temperature) and data from Scudder grid and net sweeping were collected simultaneously at Dairy A during weeks 19. These data were subjected to correlation analysis (PROC CORR, SAS 2002). Enumeration of both aerobic bacteri a and of E. coli O157:H7 included calculations of t he range, mean and median values using spreadsheet functions (Excel 2003). The number of colony forming units of plates containing 14300 CFU/plate was multiplied by the dilution factor, and the count was recorded in scientific notation to one significant digit. Minimum and maximum values of individual sample types on different days were not examined statistically. Prevalence rates of E. coli O157:H7, i.e., the percent of tested samples that tested positi ve with at least one positive isolate, were calculated for presumptive identification on both CHROMAgar and CT SMAC agar plates using spreadsheet functions (Excel 2003) Similarly, PCR c onfirmation of prevalence rates of presumptive positive samples from b oth direct culture agars was calculated using spreadsheet functions (Excel 2003). Prevalence rates were calculated by dividing the number of presumptive positive samples by the total number of tested samples, and multiplying by 100. Active fly monitoring methods were examined for relationship trends to Escherichia coli O157:H7 prevalence rates on CHROMAgar using spreadsheet functions (Excel 2003). CHROMAgar prevalence rates were used rather than PCR prevalence rates

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154 because direct culture methods were per formed on fresh samples while PCR was performed on presumptive positive isolates that had been stored for approximately 1.5 yr. Also, because only positive isolates of samples which were presumptive positive using direct culture methods were submitted to P CR, the number of samples submitted to PCR was less than the original number of samples that were examined microbiologically Data which suggested the existence of trends were then analyzed for correlations statistically using SAS with Pearsons coefficien t, as appropriate (PROC CORR, SAS 2002). Results Fly Monitoring There were no correlations at Dairy A between alsynite traps and spot cards or between alsynite traps and sticky cards in either the threeway or paired analyses during the nine weeks that al l three devices were used at Dairy A. P ositive correlation (r=0.67444, p = 0.0463) (r2 = 0.4548) between spot cards and sticky cards w as observed when data from 30 October 2008 to 4 December 2008 were subjected to threeway analysis A positive but weaker correlation (r=0.42943, p<0.0001) (r2 = 0.1844) was observed for paired analysis of card data from 23 September 2008 to 4 December 2008. During the same nine weeks at Dairy B, there was a positive correlation (r=0.69895, p=0.0362) between data from alsynit e traps and spot cards. Sticky cards were not used at Dairy B. There was a correlation of spot card numbers between dairies when all spot cards, including cards from the milk barn, were used in the analysis (F=16.06; df=1,418; P <0.001). There was stronger correlation between the spot card counts at different dairies when Dairy A's milk barn data were excluded from the analysis (F= 55.58; df=1,323; P <0.0001).

PAGE 155

155 Sweep net fly counts were higher than fly counts on Scudder grids but there was no correlation between the two active fly monitoring methods even when data were logarithmically transformed Scudder grid counts and sweep net counts were not influenced by weather conditions (barometric pressure, relative humidity, or temperature). Enumeration of Aerobi c Bacteria and Escherichia coli O157:H7 Over the entire study period from 14 June 2008 to 16 September 2008, 35 house fly, 24 spilled grain and nine manure samples were collected. Of these 68 samples, 14 fly, 17 grain and six manure samples were obtained from Dairy A while 10 fly, seven grain and thre e manure samples were obtained from Dairy B. During the same period seven and four fly samples were col le cted from Restaurants C and D, respectively. Because selective media were used for enumeration of both a erobic bacteria and E. coli O157:H7, actual counts might be underreported. Enumeration of aerobic bacteria varied across sample types and dates (Table 3 3). All sample types contained at least one sample containing 107 aerobic bacteria (Table 3 3). Mean a erobic plate counts for house flies, grain, and manure samples for the entire study period were 5.1 x106, 2.0 x 107, and 2.1 x 107 CFU/g, respectively (Table 3 4) Overall aerobic bacteria counts in grain and manure samples were 10 100 fold greater at Dair y A than at Dairy B (Table 3 4). House fly carriage of aerobic bacteria was similar in magnitude for both dairies and Restaurant C (Table 34). Flies from Dairy A had many more bacteria than flies from Dairy B (Table 3 4), which is consistent with higher g rain and manure bacterial loads at Dairy A than at Dairy B. Flies from Restaurant D had 100fold fewer bacteria than from all other sites (Table 3 4) H owever, aerobic bacter ia were enumerated from only one Restaurant D house fly sample, whereas two sample s from

PAGE 156

156 each dairy and three samples from Restaurant C were enumerated Dairy A flies averaged 8.5 x 106 aerobic bacteria and Dairy B flies averaged 1.1 x 106. Manure samples at Dairy A averaged 3.7 x 107 aerobic bacteria while manure samples from Dairy B averaged 4.6 x 106. At each dairy, aerobic bacteria counts were lower in flies than in substrate media. Dairy A grains had higher aerobic bacterial loads than manure, but Dairy B grains had lower bacterial loads than manure. Enumeration of E. coli O157:H7 for 31 May 2008 and 14 June 2008 ranged from 2.7 x 105 to 1.0 x 107 CFU/g E. coli O157:H7. These data were generated from one house fly and one manure sample on the former date and one grain and onemanure sample on the latter date Escherichia coli O157:H7 Prevalence by Direct Culture A total of 68 samples were collected from all sites, of which 57 samples were test ed using CHROMAgar plates. Collected samples consisted of 35 house fly, 24 grain, and nine manure samples. Tested samples were comprised of 33 f ly 17 grain, and seven manure samples. Time constraints prevented testing of the remaining 11 collected samples. Microbiological processing of the 57 samples produced 197 isolates that were compr ised of 103 fly, 61 grain, and 3 3 manure isolates. Of the 57 tested samples, 11 ( 19.3%) were presumptive positive for E. coli O157:H7 (Table 35 ) on CHROMAgar Across the study, E. coli O157:H7 combined sample prevalence for both dairies was 17.4% (8/ 46) with 13.0% (6/46 ) from Dairy A and 4.3% (2/ 46) from Dairy B Combined prevalence of E. coli O157:H7 at the restaurants was 27.3% ( 3/ 11) with 18.2% (2/11 ) from Restaurant C and 9.1% (1/11 ) from Restaurant D Escherichia coli O157:H7 was presumptively isolated and identified from house flies at all four sites and fro m grain at both dairies using CHROMAgar plates

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157 (Table 3 5). All manure samp les tested negative on CHROMAgar Respective overall presumptive positive prevalence rate s of E. coli O157:H7 isolated from house flies, grain, and manure samples were 1 4.0% (8/ 57), 5.3% (3 /57) and 0% (0/57). Within each tested sample type, 24.7% ( 8/33) of the flies, 1 7.6 % (3/17) of the grain, and 0. 0% (0/7) of the manure samples were presumptive positive on CHROMAgar Within samples collected only at farms, 18% (5/28) of the sample s at Dairy A and 11% (2/18) of the samples at Dai ry B were presumptive positive on CHROMAgar There was no relationship between E. coli O157:H7 presumptive positive prevalence rates and the Scudder grid fly counts or the sweep net fly counts. S imila rly, sa mples that were positive on CT SMAC agar within each tested sample type were comprised of six fl y, five grain, and two manure samples. These 13 samples tested positive on CT SMAC agar only, while nine additional samples that tested positive on CT SMAC were clones of CHROMAgar positive samples. Within samples collected only at farms, 6/9 from Dairy A 3/4 from Dairy B 2/2 from Restaurant C, and 1/2 from Restaurant D were presumptive positive on C T SMAC a gar. Escherichia coli O157:H7 Prevalence by Polymerase Chain Reaction As described previously, i solates of all 11 samples that were presumptive positive on CHROMAgar for the entire study were submitted to PCR (Table 3 5) Additionally, 13 samples that were presumptive positive on CT SMAC agar plates on 26 August 2008 and 16 September 2008 were submitted to PCR. Thus, 24 tota l samples were submitted to PCR. Twelve of 24 (50%) samples were positive using multiplex PCR. The number of samples positive for both gene fragments was increased to 14/24 (58%) when PCR was done separately for each primer pair Samples obtained in wk 6 (5 August 2008) were not available for PCR assays due to freezer thawing that killed the corresponding isolates.

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158 Specific breakdown of PCR confirmation by sample type of the samples that w ere submitted to multiplex PCR analysis revealed that 58% (7/12) of house flies, 56% (5/9) of grain and 0% (0/3) of manure samples were confirmed as positive. Additional testing by uniplex PCR increased the confirmation rates to 67% (8/12) for house flies and (1/3) for manure samples, while prevalence rates for grain were unchanged. Overall prevalence increased from 50% (12/24) to 58% (14/24). N anodrop spectrophotometry analysis resulted in 260:280 ratios that ranged from 1.82 to 1.90. The quantity of DNA template extracted from cultures ranged from 2 9 ng/l in the four isolates (31 May 2008 and 14 June 2008) that were evaluated using spectophotometry nanodrop analysis. The 16S rDNA efficiency of the 46 selected isolates was 89% (41/46 isolates). Eleven percent of the samples did not contain detectable levels of DNA This suggests that up to 11% of the 197 isolates (corresponding to the original 57 samples) that did not produce amplicons of the expected size might have contained degraded DNA. Discussion S weep net fly counts were generally higher than Scudder grid fly counts. This supports the findings of D hillon and Challet (1985), who found that fly counts with sweep nets were nearly double fly counts on Scudder grids. However, they did not report an exam ination of their data for correlation between the two fly monitoring methods. Some challenges were experienced during this study. In the field, house fly populations were highly variable throughout the sampling per iod for all monitoring methods. Thi s was particularly true for the restaurant garbage dumpsters in town, making collection of adequate numbers of flies difficult. In many instances, this may have been caused by insecticide bait or residual insecticide applications as large numbers of dead

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159 and/or twitching flies accompanied by fly bait pellets were observed at all sites on several occasions. Therefore, the microbiological protocol used for processing fly samples in this research had to be modified to accommodate collection of smaller numbers of f lies than originally planned. Decreased numbers of flies in pools did not appear to have a detrimental effect upon isolation of E. coli O157:H7, as positive fly samples were obtained continuously throughout the test period. During warmer months (summer an d fall ), sweep netted flies placed in the cooler could not be sufficiently knocked down to allow for field identification and sorting of individual flies into microcen trifuge tubes. This was likely caused by the high ambient air temperatures, which r apidly melted the ice in the cooler. As a result of these constraints, collection of flies was conducted differently during cold and hot months. Modification of the fly collection protocol may have influenced the results of this, because flies collected in warme r months remained relatively active for 2 6 h. This may have resulted in artificial or increased transfer of pathogenic microorganisms among flies held in cups that could have contributed to the high prevalence rates that I obtained from house fly samples. The use of dry ice in coolers during hot weather would lessen this effect, but dry ice production of CO2 might adversely impact aerobic growth, so that another method of implementing steady cool temperatures would be useful House fly carriage of aerobic bacteria was variable between all sites with restaurants providing counts at both extremes. At each dairy, aerobic bacteria counts were highest in manure and lowest in flies. This supports the possibility, as reported by (Vold et al. 2000) that high background counts of competing microorganisms decreases detection of E. coli O157:H7 In particular, because isolation of E. coli O157:H7 from

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160 dairy cattle manure was so low in comparison to that from house flies and spilled grain, the relationship of aerobic bacteria with E. coli O157:H7 needs to be examined further, within the various media that were tested in this experiment. Additionally, the role of house flies as a potential reservoir, not just as pathogen vectors, needs to be studied more. Enumeration of E. coli O157:H7 ranged from 2.7 x 105to 2.4 x 106 for the two samples obtained on 31 May 2008 (one house fly and one manure) and the two samples obtained on 14 June 2008 (one grain and one manure). On 31 May 2008, the house fly sample from Dairy A contain ed 2.4 x 106 CFU/g. These data are in agreement with E. coli O157:H7 and aerobic bacteria counts reported previously from house flies (Alam and Zurek 2004, Sanderson et al. 2005), cattle feed (Ahmad et al. 2007), and cattle feces (Brichta Harhay et al. 2007). The main purpose of this study was to determine prevalence rates of E. coli O157:H7 at two dairy farms and in two restaurant garbage dumpsters in a nearby town. Escherichia coli O157:H7 was isolated from house flies at all four locations, and from gra in at both dairies. Although no E. coli O157:H7 was isolated using direct culture methods from manure, only seven manure samples were tested, whereas 17 grain and 33 house fly samples were tested. Lahti et al. (2003) and Omisakin et al. (2003) observed tha t detection was directly linked to numbers of samples processed. If target pathogen numbers are low, then detection can be difficult without increased numbers of samples (Brichta Harhay et al. 2007). Thus, it is possible that E. coli O157:H7 might have bee n detected in manure if more samples had been processed. Conversely, recovery of E. coli O157:H7 from manure can be inconsistent due to high densities of competing background microorganisms (Pao et al. 2005), so that increasing the number of samples

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161 might have had no impact on prevalence rates. Zero or very low recovery rates are not uncommon in cattle feces. Hancock et al. (1994) recovered E. coli O157:H7 from only 0.28% (10 of 3,570) of dairy cattle feces samples and Galland et al. (2001) isolated E. coli O157:H7 from only 0.26% (45/17,050) of cattle fecal pats. Overall prevalence of E. coli O157:H7 at both dairies combined ranged from 3.6 10.7%, while prevalence at the restaurants combined ranged from 1.8 7.1 %. These data are in agreement with isolat ion rates obtained at dairies in some studies (Heuvelink et al. 1998, Bonardi et al. 2001, Smith et al. 2005, Oporto et al. 2008), although not as high as reported by others (Sanderson et al. 2006). The results obtained in this study with respective overa ll prevalence rates for house flies, grain, and manure samples at 1 4.0% (8/57 samples), 5.3% (3/57), and 0.0% (0/57) support the findings of previous authors (Lahti et al. 2003, Pao et al. 2003, Pearce et al. 2004, Brichta Harhay et al. 2007) that increase d sampling may be an important factor in pathogen detection. Direct culture isolation of E. coli O157:H7 from house flies was approximately 2.6 times greater than from grain. This suggests that detection of E. coli O157:H7 on dairies might be more accurate ly determined by testing house flies instead of grain or manure samples, regardless of which isolation method is utilized. In addition to providing higher prevalence of E. coli O157:H7, house flies can carry and e xcrete this pathogen for up to 4 d (Sasaki et al. 2000). Additionally, house flies can disperse from dairies to restaurants and other sites in town up to 3.0 km distant (Chapter 2). Recovery of 24.6% of E coli O157:H7 from house flies is a much higher recovery rate than those reported by Agui et al. (2001) (7.2%) and Keen et al. (2006)

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162 (5.2%), but less than that that reported by Fotedar et al. (1992) (31.233.8%). There were substantial differences in the culture and isolation methods used by Augie et al. (2001) and Keen et al. (2006) versus those methods used by Fotedar et al. (1992) and myself. Agui et al. (2001) and Keen et al. (2006) used enrichment broths that were selective for E. coli They added multiple antibiotics to the selective enrichment broths and also to the CHROMAgar plates. In con trast, both Foetedar et al. (2006) and I used a nonselective enrichment broth, to which only one antibiotic instead of multiple antibiotics was added. Also, I did not supplement the CHR OMAgar plates with antibiotics. This suggests that antibiotics might be inhibitory for isolation of E. coli O157:H7 from house flies. Thus, testing house fly samples instead of grain and manure at dairies might provide a cost savings to researchers in addition to higher prevalence rate data, because expenses associated wit h purchasing, preparing, storing, and disposing of antibiotics can be greatly reduced. Elimination of antibiotics from broths and agars would extend the shelf life of these products further increasing cost savings due to reduced labor expenses. Although CHROMAgar O157 is very selective and specific for E. coli O157, violet colored colonies presumptive for E. coli O157:H7 tended to grow slowly in this experiment. Slow growth of presumptive colonies on CHROMAgar plates was discovered when one batch of nega tive plates remained on the bench for 24 h after removed from the i ncubator. Previously, CHROMAgar plates that had not shown positive presumptive growth were discarded after the initial 24 h incubation. After this discovery, CHROMAgar plates were retained and reexamined at both 24 and 48 h after removal from incubator. Prevalence of E. coli O157:H7 on CHROMAgar plates may have been under reported in this study because most samples were subjected to 6 18 h incubation.

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163 The final batch of CHROMAgar that was used for isolation on 16 September 2008 was excessively diluted so that the agar was difficult to streak. Colonies did not remain on top, but sank into the media. Because the typical morphology and color was not discernible, all CHROMAgar plates on 16 Sept ember 2008 were recorded as negative. However, up to five isolates were transferred onto CT SMAC plates by digging a loop into "submerged" CHROMAgar colonies and streaking as usual. Seventeen of 75 isolates transferred from the "negative" CHROMAgar plates to CT SMAC plates tested positive on CT SMAC. This clearly suggests that the colonies may have been positive on the CHROMAgar media, despite the improperly made media that obscured normal reading of plates The 17 positive isolates were distributed from th e 10 different samples, so that all samples tested positive in CT SMAC with at least one positive isolate. However, all samples for this date were recorded as negative on CHROMAgar for this study (Tables 35). It is also possible that competing organisms grew on the CHROMAgar in sufficient quantities to outcompete E. coli O157:H7. Although designed for selective growth of E. coli O157:H7, closely related coliforms including Proteus spp. can grow. False positives are also a possibility on CHROMAgar that is not supplemented with potassium tellurite, because nonE. coli O157:H7 bacteria such as Salmonella spp. have the same colony color on this media (CHROMAgar protocol, Invitrogen, CA). More closely related results between CHROMAgar and CT SMAC plates might have been obtained if the CHROMAgar had been supplemented with potassium tellurite. However, this study has higher prevalence rates than studies that did supplement CHROMAgar

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164 with antibiotics, so that the impact of antibiotics upon isolation of E. coli O15 7:H7 has yet to be fully understood. One of the original goals of this study was to determine if there were correlations between fly population densities and E. coli O157:H7 prevalence using direct culture methods at Dairy A. However, due to changes in f arm management practices on Dairy A, this became a minor research topic. Specifically, the location for feeding the cattle was changed approximately halfway through the study from the milk barn to the feed barn. Therefore, only active fly monitoring methods could be examined for potential correlations to E. coli O157:H7 prevalence, because Scudder grids and sweep netting were consistently conducted at Dairy As feed barn, the microbiological specimen collection site. In contrast to active fly monitoring methods, it was not possible to look for correlations of passive fly monitoring methods to E. coli O157:H7 prevalence because spot cards and sticky cards were not in use during the entire m icrobiological sampling period. A nother goal of this study was to use PCR assays to confirm direct culture presumptive isolation and identification of E. coli O157:H7 colonies conducted in this project. In this study, only 58% (14/24) of all samples that were presumptive positive using direct culture media were confirmed by PCR analysis using both multiplex and uniplex assays despite using multiple isolates for samples. The 14 samples that were confirmed as positive by PCR comprise only 25% of the initial 57 samples that were cultured. Confirmation by PCR of only 58% of the samples suggests that the remaining 42% were either false positives on culture plates or that the fresh cultures lacked adequate

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165 quantities of DNA for successful PCR amplification following the extended time in the freezer It is interesting that 33% (1/3) of manure tested positive by uniplex PCR, because all three manure samples that were tested by PCR were obtained from isolates that appeared negative using CHROMAgar media. These results appear to confirm either an increased sensitivity of PCR over dire ct culture (Fratamico et al. 2005) or detection of sorbitolfermenting strains (Cebula et al. 1995). As with direct culture methods, PCR testing of house flies in this study provided the best method for detecting E. coli O157:H7 on dairies, and provided more information than grain or manure samples did about the presence of E. coli O157:H7 on the dairies. Therefore, house flies should be an important part of any sampling program at dairies when looking for this pathogen. Confirmation by PCR might have been reduced in this study because purified i solates were stored long term at 20 C so that thawing and refreezing of cultures might have contributed to mortality (Mennigmann 1979), gene loss (Ach et al. 2005) and/or contamination by laboratory microorgani sms ( V. Mai, personal communication) In this study, a n 8.3% mortality rate was observed over 1.5 yr which is similar to 7% mortality over nine mo reported by Doyle and Schoeni (1984) Mortality of individual isolates likely impacted the PCR confirmation rate and underscores the importance of obtaining multiple isolates from each presumptive positive colony. Survival of cultures in 20 C storage was recently shown to be increased by restricting the exponential growth phase to approximately 3 h or increas ing the nutrient availability prior to storage (Sezonov et al. 2007) Cultures in this study were provided with minimal nutrients and were incubated

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166 for 242 h, so that they might have been dead or stressed and nonviable prior to their placement in the freezer (Sezonov et al. 2007) Samples positive for fliCH7 gene segments could include E. coli of serotypes other than O157 (Cebula et al. 1995, Mead and Griffin 1998, Szalanski et al. 2004) Because the combination of fliCH7 and rfbEO157 is unique to E. coli O157:H7 (Bilge et al. 1996), a multiplex PCR assay including these two fragments was desired. I obtained only one band in several isola tes using multiplex PCR. For samples that showed only one band, I subsequently performed uniplex PCR for each gene fragment separately. In this manner, I detected the presence of genes in isolates that had previously not produc ed a band that was intense en ough to be visualized on the multiplex agarose gels. The lack of gene fragment amplification in the multiplex PCR could have been due to the depletion of nucleotides by the other primer, particularly if the primer pair for one gene was working more efficie ntly than the other pair. It was difficult to establish a suitable annealing temperature that worked well for both gene fragments. Initial attempts at multiplex PCR were conducted without success using the protocol with an annealing temperature of 48 C published by Szalanski et al. (2004). As soon as the annealing temperature was increased to desired gene fragments were successfully amplified during PCR assays. Bacterial 16S rDNA was detected by PCR for 41 of 46 (89%) samples verifying that the gene fragments obtained in the multiplex and uniplex PCR assays were due to bacterial DNA, because rDNA, located on ribosomal genes, contains nucleotide sequences that are highly conserved in all bacteria species. However, five samples (11%) tested negati ve for 16S rDNA. This can occur if the samples are contaminated with

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167 proteins or if DNA concentrations are low or sample inhibitors were not removed during DNA purification. Because DNA absorbs ultraviolet (UV) light at 260 nm and protein absorbs UV light at 280 nm, pure DNA samples will exhibit a 260:280 ratio of 1.8 2.0 (Altshuler 2006), while protein contaminated samples exhibit 260:280 ratios lower than 1.8 (Altshuler 2006). The 260:280 ratios for the four samples processed with this technique ranged from 1.82 to 1.90, which indicated that thi s representative selection of samples was relatively free from protein contamination. Although relatively free from protein contamination, the concentrations of DNA after extraction ranged from 2to 9ng/l, whi ch is low. PCR testing of fecal specimens can be very difficult, because many PCR inhibiting substances are extracted from samples along with the target DNA (Holland et al. 2000). Inhibiting factors commonly found in fecal samples include bile salts, heme, bilirubins, and complex carbohydrates (Holland et al. 2000). Because my samples were cultured on sensitive media supplemented with antibiotics, the impact of PCR inhibiting factors in the PCR assays should have been minim al All manure samples that were submitted to PCR analysis were culture negative, indicating that perhaps fecal background did inhibit direct culture detection. In the current study, PCR was able to detect one of those samples as positive, due to the combined presence of both target gene f ragments. This illustrates the sensitivity of the PCR over direct culture, and emphasizes the importance of performing both techniques. Results of PCR analysis of samples in this study cannot be directly contrasted with directculture analysis because the two methods were not used s imultaneously on fresh samples.

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168 To my knowledge t his is the first study where attempts were made to correlate house fly bacteria loads to house fly population dynamics on dairies E numeration of E. coli O157:H7 in this study wa s insufficient to fully explore this relationship but future studies of this suspected correlation could provide a valuable public health and veterinary health tool. If a correlation exists between house fly populations and pathogen prevalence, then the e pidemiological role of the house fly in a disease transmission cycle would be better understood, and possibly better quantified in terms of economic and health costs. This would emphasize the importance of having a strong IPM program for flies Additionall y, detection of E. coli O157:H7 at dairies and other livestockrearing facilities will continue to be important, particularly as human urban and suburban residential areas continue to expand. Flies provided the most reliable source of E. coli O157:H7 from dairies, and f uture research should explore direct detection of E. coli O157:H7 from house flies to better understand the role that the house fly might have in dissemination of this pathogen. Finally, the interactions of aerobic bacteria and E. coli O157:H 7 within different media on a dairy farm need to be examined more, to understand why E. coli O157:H7 is more prevalent in house flies than in dairy cattle manure.

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169 Figure 31. Scudder grid (45 x 45 cm) used to assess house fly populations on dairy farm s. Scudder grids were placed on top of spilled grains on the ground or on top of feed troughs. Fly counts consisted of the numbers of house flies that were resting on the grid five seconds after placement. Up to 10 Scudder grid counts were performed at each site. Scudder grids were placed in the same location at approximately the same time of day by the same operator to minimize variability.

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170 Figure 32. Spot cards at Dairy A's milk barn and at Dairy B were inserted into metal frames wh ich were nailed to horizontal wooden support beams 23 m above the ground. Cards were held in place by a small metal binder clip placed at the top edge of the metal frame.

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171 Figure 33. Spot and sticky card s. Spot card clipped in place by binder clips at Dairy A's feed barn at a height of 2 m above the ground. Sticky card clipped in place with two paper clips 1 cm below the spot card.

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172 A B C Figure 34. Sam pling methods for each type of collected sample. A) 100 g of grain or manure was placed in specimen cups. B) Two swabs were pre moistened with Buffered Peptone Water (BPW), exposed to grain or manure and placed in 9 ml BPW in a centrifuge tube. C) House flies that were captured using sweep nets were transferred into specimen cups.

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173 Table 3 1. D ates and house fly monitoring methods used at two Florida dairies Date Dairy A 1 Dairy B 7 June 2008 to 23 September 2008 Sweep nets, (FB) Sweep nets Scudder grids, (FB) Scudder grids Spot cards, (MB) Spot cards 23 September 2008 to 4 December 2008 S pot cards, (FB) Spot cards Alsynite traps Alsynite traps 30 October 2008 to 4 December 2008 Sticky cards, (FB) N/A 14 May 2009 to 25 June 2009 Alsynite traps Alsynite traps 1Monitoring on Dairy A was conducted at both the milk barn (MB) and the feed barn (FB). Due to discontinued use of the milk barn by the producer, the milk barn was eliminated from this study after 23 September 2008.

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174 Table 3 2. Primer nucleotide sequences used to amplify target genes in PCR assay Expected PCR Target Primer1 Se quence (5' 3') amplicon (bp) gene Specificity Citation2 Assay 1 (multiplex: O157 somatic and H7 flagellar antigens) fliCH7F GCG CTG TCG AGT TCT ATC GAG C 625 flic H7 flagellar gene 1 fliCH7R CAA CGG TGA CTT TAT CGC CCA TTC C 1 rfbEO157F CGG ACA TCC ATG TGA TAT GG 259 rfb nt 393651 of rfbO157:H7 2 rfbEO157R TTG CCT ATG TAC AGC TAA TCC 2 Assay 2 (uniplex: O157 somatic antigen) rfbEO157F CGG ACA TCC ATG TGA TAT GG 259 rfb nt 393651 of rfbO157:H7 2 rfbEO157R TTG CCT ATG TAC AGC TAA TCC 2 Assay 3 (uniplex: H7 flagellar antigen) fliCH7F GCG CTG TCG AGT TCT ATC GAG C 625 flic H7 flagellar gene 1 fliCH7R CAA CGG TGA CTT TAT CGC CCA TTC C 1 1 F, forward primer; R, reverse primer. 2 Reference: (1) (Gannon et al. 1997) ; (2) (Paton and Paton 1998)

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175 Table 3 3. Enumeration of aerobic bacteria (CFU/g) using Petrifilm Aerobic Plate Count plates inoculated with 1 ml of the unenriched sample. Aerobic Plate Count Date Site, Source Range (CFU/g) Mean (CFU/g) Median (CFU/g) 5/31/08 A, Manure a 2.5 x 10 6 2.5 x 10 6 2.5 x 10 6 6/14/08 A, Grain 9.8 x 10 7 9.8 x 10 7 9.8 x 10 7 A, Manure 2.8 x 10 7 4.6 x 10 7 3.7 x 10 7 3.7 x 10 7 B, Grain 4.7 x 10 6 4.7 x 10 6 4.7 x 10 6 B, Manure 2.5 x 10 6 6.7 x 10 6 4.6 x10 6 4.6 x10 6 C, House fly 1.3 x 10 7 3.0 x 10 7 2.2 x 10 7 2.2 x 10 7 6/23/08 A, Grain 1.6 x 10 5 2.8 x 10 6 1.5 x 10 6 1.5 x 10 6 A, House fly 1.4 x 10 7 2.0 x 10 7 1.7 x 10 7 1.7 x 10 7 B, Grain 1.8 x 10 5 3.0 x 10 5 2.4 x 10 5 2.4 x 10 5 B, House fly 2.2 x 10 5 4.0 x 10 6 2.1 x 10 6 2.1 x 10 6 C, House fly 1.3 x 10 3 1.3 x 10 3 1.3 x 10 3 8/26/08 A, Grain 2 4.8 x10 6 8.2 x 10 7 4.3 x 10 7 4.3 x 10 7 A, House fly b 3.4 x 10 4 1.5 x 10 5 9.2 x 10 4 9.2 x 10 4 B, Grain 2.9 x 10 6 3.5 x 10 6 3.2 x 10 6 3.2 x 10 6 B, House fly 1.1 x 10 5 1.4 x 10 5 1.3 x10 5 1.3 x10 5 C, House fly 1.5 x 10 4 1.5 x 10 4 1.5 x 10 4 D, House fly 2.1 x10 4 2.1 x 10 4 2.1 x 10 4 Samples were collected from two dairies and dumpsters at two restaurants in north central Florida. A=Dairy A, B=Dairy B, C=Restaurant C and D=Restaurant D. a Sample s were enumerated for aerobic plate counts, but not tested for E. coli O157:H7. b Two samples tested. Remaining data are CFU/g per plate for single samples.

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176 Table 3 4. Mean enumeration, by site and by sample type, of aerobic bacteria (CFU/g) using Petrifi lm Aerobic Plate Count plates inoculated with 1 ml of unenriched samples enumerated from collections between 31 May 2008 to 26 August 2008 on dairies and at restaurants in north central Florida Mean Aerobic Plate Count (CFU/g) Substrate Dairy A Dairy B Dairies a Restaurant C Restaurant D Restaurants a All flies a Grain 3.8 X 10 7 2.3 X 10 6 2.5 x 10 7 NA NA NA NA Manure 3.7 X 10 7 4.6 X 10 6 2.1 x 10 7 NA NA NA NA House fly 8.5 X 10 6 1.1 X 10 6 4.8 x 10 6 1.1 x 10 7 2.1 x 10 4 5.4 x 10 6 5.1 x 10 6 a Means for dai ries and restaurants were calculated by adding the means of Dairy A and Dairy B, or of Restaurant C and Restaurant D, respectively, and dividing the result by two. Means for all flies were calculated by dividing the summed means of all four sites and divid ing by four. Thus, means for dairies, restaurants and all flies were simple, not weighted. NA, Not applicable.

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177 Table 3 5. Prevalence (%) and number of E. coli O157:H7 CHROMAgar positive samples and number of CT SMAC and PCRpositive samples relative to th e number of samples tested after collection from two dairies and two restaurant dumpsters in north central Florida CHROMAgar CT SMAC PCR Date Site Sample Type No. Collected No. Tested No. Positivea ( %) No. Tested No. Positive No. Tested No. Positive 6/14/2008 A Grain 2 1 1 (100) 1 1 Manure 2 1 0 (0) 1 0 B Grain 2 1 0 (0) 1 0 Manure 2 1 0 (100) 1 1 C Fly 1 1 0 (0) 1 1 6/23/2008 A Fly 1 1 0 (0) NA NA NA NA Grain 4 1 0 (0) NA NA NA NA B Fly 1 1 0 (0) NA NA NA NA Grain 1 1 0 (0) NA NA NA NA C Fly 1 1 0 (0) NA NA NA NA 7/20/2008 A Fly 3 2 1 (50) 1 1 B Fly 3 2 0 (0) 0 NA C Fly 1 1 1 (100) 1 1 7/28/2008 A Fly 1 1 0 (0) NA NA NA NA B Fly 2 2 0 (0) NA NA NA NA C Fly 1 1 0 (0) NA NA NA NA D Fly 1 1 0 (0) NA NA N A NA 8/5/2008 A Fly 3 3 1 (33) NA NA NA NA Grain 3 1 0 (0) NA NA NA NA Manure 2 2 0 (0) NA NA NA NA B Fly 2 2 0 (0) NA NA NA NA Grain 1 1 0 (0) NA NA NA NA

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178 CHROMAgar CT SMAC PCR Date Site Sample Type No. Collected No. Tested No. Positivea ( %) No. Tested No. Positive No. Tested No. Positive Manure 1 1 0 (0) NA NA NA NA C Fly 1 1 1 (100) NA NA NA NA D Fly 1 1 0 (0) NA NA NA NA 8/26/2008 A Fly 5 5 2 (40) 5 2 2 1 Grain 6 6 1 (17) 6 2 5 3 B Fly 1 1 1 (100) 1 0 1 1 Grain 1 1 1 (100) 1 1 1 1 C Fly 1 1 0 (0) 1 1 1 1 D Fly 1 1 1 (100) 1 1 1 1 9/16/2008 A Fly 1 1 0 (0) 1 1 1 1 Grain 2 2 0 (0) 2 2 1 1 Manure 2 2 0 (0) 2 2 1 1 B Fly 1 1 0 (0) 1 1 1 1 Grain 2 2 0 (0) 2 2 2 0 C Fly 1 1 0 (0) 1 1 1 1 D Fly 1 1 0 (0) 1 1 1 1 TOTAL 68 57 11 (19.2) 32 22 24 14 Sites: A, Dairy A; B, Dairy B; C, dumpster at Restaurant C; D, dumpster at Restauran t D. Spilled rain and manure samples consisted of either 25 g substrate or swabs. Fly samples consisted of pools of up to 25 house flies. Distance (km) from release site (Chapter 2) is also provided. CT SMAC, sorbitol MacConkey agar supplemented with cefixime and potas sium tellurite. a Percent positive was calculated by dividing the number of positive samples by the number of samples tested in the laboratory.

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179 CHAPTER 4 OVERALL CONCLUSIONS Conclusions The overall goal of this research was to determine the role of house flies, Musca domestica L., in the transmission of Escherichia coli O157:H7 at the rural urban interface using two dairies and a small town. This was accomplished through three sampling parameters: by examin in g house fly dispersal, house fly population patt ern s, and Escherichia coli O157:H7 prevalence. S pecifically, studies were conducted to determine if house flies disperse from dairies into town, what their populations on the farms were at the time of the dispersal studies, and characterizing E. coli O157: H7 prevalence from house flies at both the dairies and in town using CHROMAgar and sorbitol MacConkey agar (CT SMAC) direct culture methods. Additionally, a rapid multiplex polymerase chain reaction (PCR) method was developed to detect the rfbEH7 and fliCO157 virulence factor genes of E. coli O157:H7 isolated from house flies, grain and manure. Escherichia coli O157:H7 prevalence data was used in conjunction with house fly dispersal data from the dairies to the town to estimate the public health risk that house flies may present in regards to transmission of E. coli O157:H7 from dairies into towns located in northcentral Florida. Background Previous research documented E. coli O157:H7 isolation from house flies on dairies (Zhao et al. 1995) and at restaur ants (Butler et al. 2010). Laboratory studies have shown that E. coli O157:H7 survives and is excreted in a viable, infectious state for up to 4 d after it is ingested by house flies (Sasaki et al. 2000) while Petridis et al. ( 2006) and Macovei et al. ( 2008) demonstrated replication of E. coli O157:H7 in house flies Field studies have shown that house flies can disperse up to 4.8 km ( Quarterman et al. 1954, Shura Bura et al. 1962) and that flies are

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180 capable of flying at rates of 1 km/h (Shura Bura et al. 1962) However, no studies have been conducted to investigate the synergism of house fly dispersal and E. coli O157:H7 survival in the natural environment. Successful isolation of this pathogen from flies located both at the dairies and in town followed by genetic analysis of both the flies and the bacteria would provide more information about origin of bacterial strains isolated from flies. Isolation by direct culture reveals important information about the biochemical characteristics of bacteria, but is olation by DNA based methods such as PCR permit serotyping and phylogenetic analysis. In addition, PCR is more sensitive and specific than direct culture techniques Both direct culture and PCR methods have been used to isolate E. coli O157:H7 from house f lies, grain, and manure samples on dairies. Pulsed field gel electrophoresis is the preferred standard for DNA based i dentification methods, but it is more time consuming and labor intensive than PCR. Conclusions I hypothesize that the house fly provides a mobile element of disease causing pathogen transmission from dairy cattle to humans, particularly in locations where the rural urban interface is within the typical flight range of house flies. Furthermore, I hypothesize that the role of house fly pathogen transmission can be better understood by genetic serotyping of both house fly and bacteria and determining if the same strains exist in both the rural and urban locations where fly transmission is suspected. I hypothesize that PCR can be used to quickl y and efficiently confirm E. coli O157:H7 isolation from house flies, grain and manure. This study provide d evidence of house fly dispersal from a dairy into town across the rural urban interface, using roads as landscape corridors, and by direct flight overland Furthermore, dispersal from a dairy to a restaurant was observed. This study demonstrate d the

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181 usefulness of a newly designed multiplex PCR assay for confirmation of direct culture presumptive positive E. coli O157:H7 samples. Future Research Ther e are additional studies that need to be conducted to build on the findings of this research in order to better understand, and perhaps eventually quantify, the role of house flies in pathogen transmission across the rural urban interface from dairies into town. In particular, more efficient DNA based isolation methods are needed. In depth understanding of house fly bacterial loads has been limited by the need to use fresh samples containing viable bacteria for direct culture isolation. Isolation of bacteria by direct culture permits long term storage of viable isolates so that DNA based analysis can be conducted at future dates. Therefore, direct culture methods are valuable and should be used. However, PCR can amplify both viable and non viable bacteria, so that storage of flies might avoid the need to culture bacteria samples. This could reduce expenses and labor, as well as increase specificity and selectivity. Future research should be conducted to determine if PCR can rapidly and efficiently isolate E coli O157:H7 directly from house fly samples that have been in longterm storage at 20 C. The relationship between house flies and E. coli O157 :H7 and other enteric bacteria should be examined for specific geographical regions by using PCR on flies tha t have been in longterm storage. Use of flies that have been in long term storage permits more extensive sampling during seasons of dominant fly activity because PCR assays can be conducted outside of the fly season during periods of time when researchers have greater available labor Summary In summary, this research has led to a better understanding of the house fly role in transmission of E. coli O157:H7 from dairies to urban areas. This research has resulted in the

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182 development of a multiplex PCR assa y for identifying E. coli O157:H7 from house flies. Results from this dissertation demonstrate that house flies provide valuable information regarding a mobile element for pathogen transmission that is lacking in grain and manure samples. House flies provi de a readily available, high return sampling option and should be incorporated in the design of a pathogen monitoring program s on dairies.

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183 LIST OF REFERENCES Abe, K., S. Yamamoto, and K. Shinagawa. 2002. Economic impact of an Escherichia coli O157:H7 out break in Japan. J. Food Prot. 65: 6672. Ach, S. J., I. Khn, G. Mbazima, P. Colque Navarro, and R. Mllby. 2005. Changes of viability and composition of the Escherichia coli flora in faecal samples during long time storage. J. Microbiol. Meth. 63: 229238. Agui, N. 2001. Flies carrying enterohemorrhagic Escherichia coli (EHEC) O157 in Japan: a nationwide survey. Med. Entomol. Zool. 52: 97103. Ahmad, A., T. G. Nagaraja, and L. Zurek. 2007. Transmission of Escherichia coli O157:H7 to cattle by house fli es. Prev. Vet. Med. 80: 74 81. Alam, M. J. and L. Zurek. 2004. Association of Escherichia coli O157:H7 with houseflies on a cattle farm. Appl. Environ. Microbiol. 70: 7578 7580. Alam, M. J. and L. Zurek. 2006. Seasonal prevalence of Escherichia coli O157:H7 in beef cattle feces. J. Food Prot. 69: 30183020. Ananth, G. P., D. C. Bronson, and J. K. Brown. 1992. Generation of airborne fly body particles by four electrocution fly traps and an electronic fly trap. Inter. J. Environ. Health Res. 2: 106113. A nderson, G. S. and B. J. Danielson. 1997. The effects of landscape composition and physiognomy on metapopulation size: the role of corridors. Landscape Ecol. 12: 261271. Anderson, J. and J. Poorbaugh. 1964. Biological control possibility for house flies. Cal. Agr. 18: 24. Angelo, M. J. and F. Slansky. 1984. Body building by insects: trade offs in resource allocation with particular reference to migratory species Fl a Entomol. 67: 2241. ANON. 1940. Summer diarrhea and horsedrawn vehicles. Am. J. Pub. H ealth 30: 825 826. Archer, D. L. and F. E. Young. 1988. Contemporary issues: Diseases with a food vector. Clin. Microbiol. Rev. 1: 377398. Asakura, H., S. Makino, T. Shirahata, T. Tsukamoto, H. Kurazono, T. Ikeda, and K. Takeshi. 1998. Detection and genetical characterization of Shiga toxin producing Escherichia coli from wild deer. Microbiol. Immunol. 42: 815822. Avancini Rita MP and A. R. Silveira Gerson. 2000. Age structure and abundance in populations of muscoid flies from a poultry facility in Southeast Brazil. Mem. Inst. Oswaldo Cruz.95: 259264. ( http://www.scielo.br/scielo.php?script=sci_arttext&pid =S007402762000000200022&lng=en)

PAGE 184

184 Avery, L. M., A. P. Williams, K. Killham, and D. L. Jones. 2008. Survival of Escherichia coli, O157:H7 in waters from lakes, rivers, puddles and animal drinking troughs. Sci. Total Environ. 389: 378385. Axtell, R. C. 1970. Integrated fly control program for cag ed poultry houses. J. Econ. Entomol. 63: 400405. Bailey, D. L. 1970. Forced air for separating pupae of house flies from rearing medium. J. Econ. Entomol. 63: 331333. Baldwin, T. and E. H. Bryant. 1981. Effect of size upon mating performance within geographic strains of the housefly, Musca domestica L. Evol. 35: 11341141. BarkocyGallagher, G. A., K. K. Edwards, X. Nou, J. M. Bosilevac, T. M. Arthur, S. D. Shackelford, and M. Koohmaraie. 2005. Methods for recovering Escherichia coli O157:H7 from catt le fecal, hide, and carcass samples: sensitivity and improvements. J. Food Prot. 68: 22642268. Barnard, D. R. 2003. Control of fly borne diseases. Pestic. Outlook 14: 222228. Bell, B. P., M. Goldoft, P. M. Griffin, M. A. Davis, D. C. Gordon, P. I. Tarr C. A. Bartleson, J. H. Lewis, T. J. Barrett, J. G. Wells, et al. 1994. A multistate outbreak of Escherichia coli O157:H7associated bloody diarrhea and hemolytic uremic syndrome from hamburgers : the Washington experience. J. Am. Med. Assoc. 272: 13491353. Berg, H. C. 2004. E. coli in motion (biological and medical physics: biomedical engineering). AIP Press, Springer Verlag, New York, NY. Berry, I. L., P. J. Scholl, and J. I. Shugart. 1981. A mark and recapture procedure for estimating population sizes of stable flies. Environ. Entomol. 10: 8893. Bertani, G. 1951. Studies on lysogenesis. I. The mode of phage liberation by lysogenic Escherichia coli J. Bacteriol. 62: 293300. Bertani, G. 2004. Lysogeny at midtwentieth century: P1, P2, and other experimental systems. J. Bacteriol. 186: 595600. Bettelheim, K. A. 1991. The genus Escherichia pp. 2696 2736. In A. Balows, H. G. Tr per, M. Dworkin, W. Har der, and K.H. Schleifer [eds.], The prokaryotes: a handbook on the biology of bacteria: ecophysiology, isolation, identification, applications. Springer Verlag, New York, NY. Beutin, L., G. Krause, S. Zimmermann, S. Kaulfuss, and K. Gleier. 2004. Characterization of Shiga toxin producing Escherichia coli strains isolated from human patients in Germany over a 3year period. J. Clin. Microbiol. 42: 10991108.

PAGE 185

185 Beutin, L., A. Miko, G. Krause, K. Pries, S. Haby, K. Steege, and N. Albrecht. 2007. Identification of humanpathogenic strains of Shiga toxin producing Escherichia coli from food by a combination of serotyping and molecular typing of Shiga toxin genes. Appl. Environ. Microbiol. 73: 4769 4775. Bilge, S. S., J. C. Vary, S. F. Dowell, and P. I. Tarr. 1996. Role of the Escherichia coli O157:H7 O side chain in adherence and analysis of an rfb locus. Inf ect. Immun. 64: 47954801. Birkemoe, T. A. Soleng, and A. Aak. 2008. Biological control of Musca domestica and Stomoxys calcitrans by mass releases of the parasitoid Spalangia cameroni on two Norwegian pig farms. Biocontrol 54: 425436. Bishopp, F. C. a nd E. W. Laake. 1921. The dispersion of flies by flight. J. Agric. Res. 21: 729766. Black IV, W. C. and E. S. Krafsur. 1986. Geographic variation in house fly size adaptation or larval crowding ? Evol. 40: 204206. Black IV, W. C. and E. S. Krafsur. 2008. Fecundity and size in the housefly: investigations of some environmental sources and genetic correlates of variation. Med. Vet. Entomol. 1: 369382. [BLAST] Basic Local Alignment Search Tool. 2009. Basic local alignment search tool. ( http://blast.ncbi.nlm.nih.gov/ Blast.cgi ) Bodnaryk, R. P. and P. E. Morrison. 1966. The relationship between nutrition, haemolymph proteins, and ovarian development in Musca domestica J. Ins. Physiol. 12: 963976. Bolton D.J., Byrne C.M. Sheridan, J.J., McDowell D.A. and Blair I.S. 1999. The survival characteristics of a non toxigenic strain of Escherichia coli 0157:H7. J. Appl. Microbiol. 86: 407411. Borror, D. J., C. A. Triplehorn, and N. F. Johnson. 1989. An introduction to the study of insects, 6th ed. Saunders College Publishers, Orlando, FL Bosik, J. J. 1997. Common names of insects and related organisms. Entomological Soc iety of America, College Park, MD. Boulesteix, G., P. Le Dantec, B. Chevalier, M. Dieng, B. Niang, and B. Diatta. 2005. Role of Musca domestica in the transmission of multiresistant bacteria in the centres of intensive care setting in sub Saharan Africa. Ann. Fr. Anesth. Reanim. 24: 361365. Bracken, G. K. 1965. Effects of dietary components on fecundity of the parasitoid Exeristes comstockii (Cress) (Hymenoptera: Ichneumonidae). Can. Entomol. 97: 10371041.

PAGE 186

186 Brandl, M. T. 2006. Fitness of human enteric pathogens on plan3ts and implications for food safety. Annu. Rev. Phytopathol. 44: 367 392. Breuer, T., D. H. Benkel, R. L. Shapiro, W. N. Hall, M. M. Winnett, M. J. Linn, J. Neimann, T. J. Barrett, S. Dietrich, F. P. Downes, D. M. Toney, J. L. Pearson, H. Rolka, L. Slutsker, P. M. Griffin, and the Investigation Team. 2001. A multistate outbreak of Escherichia coli O157:H7 infections linked to alfalfa sprouts grown from contaminated seeds. Emerg. Infect. Dis. 7: 977982. Brichta Harhay, D. M., M. Arthur, J. M. Bosileva, M. N. Guerini, N. Kalchayanand, M. and Koohmaraie. 2007. Enumera tion of Salmonella and Escherichia coli O157:H7 in ground beef, cattle carcass, hide and faecal samples using direct plating methods. J. Appl. Microbiol. 103: 16571668. Broce, A. B. 1988. An improved alsynite trap for stable flies, Stomoxys calc itrans (D iptera: Muscidae). J. Med. Entomol. 25: 406409. Broce, A. B. 1993a. Dispersal of house flies and stable flies. pp. 61 69. In G. D. Thomas and S. R. Skoda [eds.]. Rural flies in the urban environment? North Central Regional Res. Bull. 335, Institute of Agriculture and Natural Resources Res. Bull. 317. University of Nebraska Agricultural Research Division, Lincoln, NE. Broce, A. B. 1993b Electrocuting and electronic insect traps: trapping efficiency and produc tion of airborne particles. Int J. Environ. H ealth Res. 3: 4758. Broce, A. B. and R. J. Elzinga. 1984. Comparison of prestomal teeth in the face fly ( Musca autumnalis ) and the house fly ( Musca domestica ) (Diptera: Muscidae). J. Med. Entomol. 21: 8285. Broce, A. B. and J. E. Urban. 1998. Potential microbial health hazards associated with operation of bug zappers. Ann. Mee ting Am. Soc. Microbiol. Q 252. Buma, R., T. Maeda, Y. Marutaka, M. Kamei, H. Nagamune, and H. Kourai. 2004. Vectorial capacity of larvae, pupae and adult of housefly ( Musca domes tica ) for Escherichia coli O157:H7 and the possibility of transmission from source to human. Med. Entomol. Zool. 55: 95106. Buma, R., H. Sanada, T. Maeda, M. Kamei, and H. Kourai. 1999. Isolation and characterization of pathogenic bacteria, including Esc herichia coli O157:H7, from flies collected at a dairy farm field. Med. Entomol. Zool. 50: 313321. Burg, J. G. and R. C. Axtell. 1984. Monitoring house fly, Musca domestica (Diptera: Muscidae), populations in caged layer poultry houses using a baited jugtrap. Environ. Entomol. 13: 10831090.

PAGE 187

187 Butler, J., A. GarciaMaruniak, F. Meek, and J. E. Maruniak. 2010. Wild Florida house flies ( Musca domestica L.) as carriers of pathogenic bacteria. Fl a Entomol. 93: 218223. Buzby, J. C., T. Roberts, C. T. Jordan Lin, and J. M. MacDonald. 1996. Bacterial foodborne disease: medical costs and productivity losses. Agr. Econ. Rep. No. AER741, August 1996. Byrd, J. H. and J. L. Castner. 2009. Forensic entomology: the utility of arthropods in legal investigations, 2nd ed., CRC Press, Boca Raton, FL. Cafarchia, C., R. P. Lia, D. Romito, and D. Otranto. 2009. Competence of the housefly, Musca domestica, as a vector of Microsporum canis under experimental conditions. Med. Vet. Entomol. 23: 2125. Cagney, C., H. Crowley, G. Duffy, J. J. Sheridan, S. O'Brien, E. Carney, W. Anderson, D. A. McDowell, I. S. Blair, and R. H. Bishop. 2004. Prevalence and numbers of Escherichia coli O157:H7 in minced beef and beef burgers from butcher shops and supermarkets in the Republic of Ir eland. Food Microbiol. 21: 203212. Callaway, T. R., R. C. Anderson, T. S. Edrington, K. J. Genovese, K. M. Bischoff, T. L. Poole, Y. S. Jung, R. B. Harvey, and D. J. Nisbet. 2004. What are we doing about Escherichia coli O157:H7 in cattle? J. Anim. Sci. 82: E93 E99. Callaway, T. R., R. O. Elder, J. E. Keen, R. C. Anderson, and D. J. Nisbet. 2003. Forage feeding to reduce preharvest Escherichia coli populations in cattle, a review. J. Dairy Sci. 86: 852860. 2005. Enterohaemorrhagic Escherichia coli : emerging issues on virulence and modes of transmission. Vet. Res. 36: 289311. Carlson, D. A., U. R. Bernier, J. A. Hogsette, and B. D. Sutton. 2001. Distinctive hydrocarbons of the black dump fly, Hydrotaea aenescens (Diptera: Muscidae). Arch. Insect Biochem. Physiol. 48: 167178. [ CDC] Centers for Disease Control and Prevention 1983. International notes: outbreak of hemorrhagic colitis -Ottawa, Canada. Mo rb. Mort. Wkly. Rep. 32: 133134. ( http://www.cdc.gov/mmwr/preview/mmwrhtml/00001271.htm ) [ CDC] Centers for Disease Control and Prevention 1994. E. coli O157:H7: procedure for isolation and identification from stool specimens. ( http://wonder.cdc.gov/wonder/ PrevGuid/p0000445/P0000445.asp). [ CDC] Centers for Disease Control and Prevention 2008. Shiga toxin producing Escherichia coli (STEC) case surveillance. ( http:// www.cdc.gov/outbreaks .html )

PAGE 188

188 [CDC] Centers for Disease Control and Prevention. 2009. Multistate outbreak of E. coli O57 :H7 infections associated with beef from JBS Swift Beef Company. ( http://www.cdc.gov/ecoli/2009/0701.html ) Cebula, T. A., W. L. Payne, and P. Feng. 1995. Simultaneous identification of strains of E scherichia coli serotype O157:H7 and their Shiga like toxin type by mismatch amplification mutation assay multiplex PCR. J. Clin. Microbiol. 33: 248 250. Chapin, J. B. 1989. Common names of insects. Bull. Entomol. Soc. Am. 35: 177180. Chapman, P. A. and C. A. Siddons. 1996. A comparison of immunomagnetic separation and direct culture for the isolation of verocytotoxin producing Escherichia coli O157 from cases of bloody diarrhoea, nonbloody diarrhoea and asymptomatic contacts. J. Med. Microbiol. 44: 267271. Chapman, P. A., C. A. Siddons, A. T. Cerdan Malo, and M. A. Harkin. 1997. A 1 year study of Escherichia coli O157 in cattle, sheep, pigs and poultry. Epidemiol. Infect. 119: 245 250. Chapman, P. A., C. A. Siddons, D. J. Wright, P. Norman, J. Fox, and E. Crick. 1993. Cattle as a possible source of verocytotoxin producing Escherichia coli O157 infections in man. Epidemiol. Infect. 111: 439447. Chapman, P. A., D. J. Wright, and C. A. Siddons. 1994. A comparison of immunomagnetic separation and direct culture for the isolation of verocytotoxin producing Escherichia coli O157 from cases bovine faeces. J. Med. Microbiol 40: 424427. Chou, C. C. and S. J. Cheng. 2000. Recovery of low temperature stressed E. coli O157:H7 and its susceptibility to crystal violet, bile salt, sodium chloride and ethanol. Int. J. Food Microbiol. 61: 127136. Christensen, C. M. 1982. External parasites of dairy cattle. J. Dairy Sci. 65: 12892193. Cirillo, V. J. 2006. Winged sponges: Houseflies as carriers of typhoid fever i n 19thand early 20thcentury military camps. Perspect. Biol. Med. 49: 5263. Cohen, D., M. Green, C. Block, R. Slepon, R. Ambar, S. S. Wasserman, and M. M. Levine. 1991. Reduction of transmission of shigellosis by control of houseflies ( Musca domestica ) Lancet. 337: 993997. Conn, D. B., J. Weaver, L. Tamang, and T. K. Craczyk. 2007. Synanthropic flies as vectors of Cryptosporidium and Giardia among livestock and dairy collected life in a multispecies agricultural complex. Vec tor Borne Zoonotic Dis. 7: 63652.

PAGE 189

189 Cornick, N. A., S. L. Booher, T. A. Casey, and H. W. Moon. 2000. Persistent colonization of sheep by Escherichia coli O157:H7 and other E. coli pathotypes. Appl. Environ. Microbiol. 66: 49264934. Couteau, D., A. L. McCartney, G. R. Gibson, G. Williamson, and C. B. Faulds. 2001. Isolation and characterization of human colonic bacteria abl e to hydrolyse chlorogenic acid J. Appl. Microbiol. 90: 873881. Crespo, D. C., R. E. Lecuona, and J. A. Hogsette. 1998. Biological control: an important component in integrated management of Musca domestica (Diptera: Muscidae) in caged layer poultry houses in Buenos Aires, Argentina. Biol. Control 13: 1624. [CSTE] Council of State and Territorial Epidemiologists. 2005. Revision of the enterohemorrhagic Escherichia coli (EHEC) condition name to Shiga toxinproducing Escherichia coli (STEC) and adoption of serotype specific national reporting for STEC. Position Statement 05ID 07. ( http ://www.Cste.Org/Position%20statements/ Searchbyyear2005) Davis, M. A. 2006. Comparison of cultures from rectoanal junction mucosal swabs and feces for detection of Escherichia coli O157 in dairy heifers. Appl. Environ. Microbiol. 72: 3766 3770. Davis, M. A., K. A. Cloud Hansen, J. Carpenter, and C. J. Hovde. 2005. Escherichia coli O157:H7 in environments of culture positive cattle. Appl. Environ. Microbiol. 71: 68166822. De Jesus, Antonio J.; Olsen, Alan R.; Bryce, John R., and Whiting, Richard C. 2004. Quantitative contamination and transfer of Escherichia coli from foods by houseflies, Musca domestica L. (Diptera: Muscidae). Int. J. Food Microbiol. 93: 259262. de la TorreBueno, J. R., S. W. Nichols, G. S. Tulloch, and R. T. Schuh. 1989. The Torre Bueno glossary of entomology. New York Entomological Society, New York, NY. Delthier I. R. 1976. The hungry fly. Harvard University Press, Cambridge, MA Denholm, I., R. M. Sawicki, and A. W. Farnham. 1985. Factors affecting resistance to insecticides i n house flies, Musca domestica L. (Diptera:Muscidae). IV. The population biology of flies on animal farms in south eastern England and its implications for the management of resistance. Bull. Entomol. Res. 1985; 75: 144158. Denny, J., M. Bhat, and K. Eck mann. 2008. Outbreak of Escherichia coli O157:H7 associated with raw milk Consumption in the Pacific Northwest. Foodborne Path. Dis. 5: 321328. Desmarchelier, P. M., S. S. Bilge, N. Fegan, L. Mills, J. C. Vary Jr ., and P. I. Tarr. 1998. A PCR specific fo r Escherichia coli O157 based on the rfb locus encoding O157 lipopolysaccharide. J. Clin. Microbiol. 36: 18011804.

PAGE 190

190 Dhillon, M. S., and G. L. Challet. 1985. The evaluation of three sampling techniques for the determination of fly (Diptera) densities at fou r sanitary landfills in southern California. Bull. Soc. Vect. Ecol. 10: 3640. Doane, C. A., P. Pangloli, H. A. Richards, J. R. Mount, D. A. Golden, and F. A. Draughon. 2002. Occurrence of Escherichia coli O157:H7 in diverse farm environments. J. Food Pro t. 70: 610. Donnenberg, M. S. 2002. Introduction. pp. xxi xxv. In Donnenberg, M. S. [ed.]. Escherichia coli : Virulence mechanisms of a versatile pathogen. Academic Press, San Diego CA. Donnenberg, M. S. and T. S. Whittam. 2002. Pathogenesis and evoluti on of virulence in enteropathogenic and enterohemorrhagic Escherichia coli J. Clin. Investigation. 107: 539548. Dougherty, E. C. 1959. Introduction of axenic culture of invertebrate metazoa: a goal. Ann. N. Y. Acad. Sci. 77: 2754. Doyle, M. E., J. Arc her, C. W. Kaspar, and R. Weiss 2006. FRI Briefings: Human illness caused by E. coli O157:H7 from food and nonfood sources. Food Res earch Inst itute, University of WisconsinMadison Madison, WI. Doyle, M. P. and L. R. Beuchat. 2007. Food microbiology: f undamentals and frontiers. ASM Press, Washington, D.C. Doyle, M. P. and J. L. Schoeni. 1984. Survival and growth characteristics of Escherichia coli associated with hemorrhagic colitis. Appl. Environ. Microbiol. 48: 855856. Duffy, G. 2003. Verocytoxigen ic Escherichia coli in animal faeces, manures and slurries. J. Appl. Microbiol 94: 94S 103S. Dunn, J. R., J. E. Keen, D. Moreland, and R. A. Thompson. 2004a. Prevalence of Escherichia coli O157:H7 in white tailed deer from Louisiana. J. Wildlife Dis. 40: 361 365. Dunn, J. R., J. E. Keen, and R. A. Thompson. 2004b. Prevalence of shigatoxigenic Escherichia coli O157:H7 in adult dairy cattle. J. Am. Vet. Med. Assoc. 224: 11511158. DuPont, H. L. 2007. The growing threat of foodborne bacterial enteropathoge ns of animal origin. Clin. Infect. Dis. 45: 13531361. Duriez, P., Y. Zhang, Z. Lu, A. Scott, and E. Topp. 2008. Loss of virulence genes in Escherichia coli populations during manure storage on a commercial swine farm. Appl. Environ. Microbiol. 74: 39353942.

PAGE 191

191 Durso, L. M. and J. E. Keen. 2007. Shiga toxigenic Escherichia coli O157 and nonShiga toxigenic E. coli O157 respond differently to culture and isolation from naturally contaminated bovine faeces. J. Appl. Microbiol. 103: 24572464. Dynal. 2007. Dynabeads anti E. coli 0157 Manual. Invitrogen Dynal AS. Oslo, Norway. Echeverria, P., B. A. Harrison, C. Tirapat, and A. McFarland. 1983. Flies as a source of enteric pathogens in a rural village in Thailand. Appl. Environ. Microbiol. 46: 3236. Echeverry, A., G. H. Loneragan, B. A. Wagner, and M. M. Brashears. 2005. Effect of intensity of fecal pat sampling on estimates of Escherichia coli O157 prevalence. A m. J V et. R es. 66: 20232027. Eddy, G. W., A. R. Roth, and F. W. Plapp, Jr. 1962. Studies on the flight habits of some marked insects. J. Econ. Entomol. 55: 603608. [ ESA] Entomological Society of America. 2009. ESA common names of insects and related organisms online database. ( http://www.entsoc.org/Pubs/Common_Names/ search.asp Accessed 11 January 2008) Euzeby, J. P. M. 2008. List of prokaryotic names with standing in nomenclature (LPSN): formerly, list of bacterial names with standing in nomenclature (LBSN). ( http://www.bacterio.cict.fr/aldl.html) [Excel] Microsoft Excel. 2003. Microsoft Excel Users Manual, Ver. 11. Microsoft Corp., Redmond, WA. [ FDACFSAN] Food and Drug Administration, Center for Food Safety and Appl ied Nutrition. 2007a. Media i ndex for BAM Bacteriological Analytical Manual (BAM) Online. ( http://www.fda.gov/Food/ScienceResearch/LaboratoryMethods/ BacteriologicalAnalyticalManualBAM/ucm055778.htm ). [ FDACFSAN] Food and Drug Administration, Center for Food Safety and Applied Nutrition. 2007b. Diarrheagenic Escherichia coli Bacteriological Analytical Manual (BAM) Online. ( http://www.cfsan.fda.gov/~ebam/bam 4a.html10 p) [ FDACFSAN] Food and Drug Administration, Center for Food Safety and Applied Nutrition. 2007c. Escherichia coli O157:H7. Foodborne pathogenic micoorganisms and natural toxins handbook: the "bad bug book." ( http://www.cfsan.fda.gov/~mow/ chap15.html ) Feder, I., F. M. Wallace, J. T. Gray P. Fratamico, P. J. FedorkaCray, R. A. Pearce, J. E. Call, R. Perrine, and J. B. Luchansky. 2003. Isolation of Escherichia coli O157:H7 from intact colon fecal samples of swine. Emerg. Infect. Dis. 9: 380383.

PAGE 192

192 Feng, P. C. H. and S. R. Monday. 2005. Multip lex PCR for specific identification of enterohemorrhagic Escherichia coli strains in the O157:H7 complex. In C. C. Adley [ed.] Methods in Biotech., Vol. 21: Foodborne pathogens: methods and protocols. Humana Press Inc, Totowa, NJ. Floate, K. D., P. Cogh lin, and G. A. P. Gibson. 2000. Dispersal of the filth fly parasitoid Muscidifurax raptorellus (Hymenoptera: Pteromalidae) following mass releases in cattle confinements. Biol. Control: 172 178. Floyd, T. M. and B. H. Cook. 1954. The housefly as a carrier of pathogenic human enteric bacteria in Cairo. J. Egypt. Publ. Health Assoc. 28: 7585. Fode Vaughn, K. A., J S. Maki, J. A. Benson, and M. L. P. Collins. 2003. Direct PCR detection of Escherichia coli O157:H7. Lett. Appl. Microbiol. 37: 239243. [ Food Consumer ] Food Consumer 2006. Nationwide E. coli O157:H7 outbreak: questions and answers. ( http://www.foodconsumer.org/777/8/Nationwide_E_Coli_O 157_H7_ Outbreak_Questions_amp_Answers.shtml ). Foster, S. P. and M. O. Harris. 1997. Behavioral manipulation methods for insect pest management. Annu. Rev. Entomol. 42: 123146. Fotedar, R., U. Banerjee, S. Singh, Shriniwas, and A. K. Verma. 1992. The h ousefly ( Musca domestica ) as a carrier of pathogenic microorganisms in a hospital environment. J. Hosp. Infect. 20: 209 215. Frandson, R. D. 1969. Anatomy and physiology of farm animals. Lea and Febiger, Philadelphia, PA. Fratamico, P. M. and D. O. Bayles 2005. Molecular approaches for detection, identification, and analysis of foodborne pathogens, pp. 114. In A. K. Bhunia, and J. L. Smith (eds.). Foodborne pathogens. Caister Acad. Press, Norfolk, UK. Fratamico, P. M., S. K. Sackitey, M. Wiedmann, and M Y. Deng. 1995. Detection of Escherichia coli O157:H7 by multiplex PCR. J. Clin. Microbiol. 33: 21882191. Fremaux, B., C. Prigent Combaret, and C. VernozyRozand. 2008. Longterm survival of Shiga toxin producing Escherichia coli in cattle effluents and environment: An updated review. Vet. Microbiol. 132: 118. Frenzen, P. D., A. Drake, and F. J. Angulo. 2005. Economic cost of illness due to Escherichia coli O157 infections in the United States. J. Food Prot. 68: 26232630. Frick, T. B. and D. W. Tallamy. 1996. Density and diversity of nontarget insects killed by suburban electric insect traps. Entomol. News 107: 7782.

PAGE 193

193 Fried, J. H., D. J. Levey, and J. A. Hogsette. 2005. Habitat corridors function as both drift fences and movement conduits for disper sing flies. Oecologia 143: 645651. Galland, J. C., D. R. Hyatt, S. S. Crupper, and D. W. Acheson. 2001. Prevalence, antibiotic susceptibility, and diversity of Escherichia coli O157:H7 isolates from a longitudinal study of beef cattle feedlots. Appl. Environ. Microbiol. 67: 16191627. Gannon, V. P. J., S. D'Souza, T. Graham, R. K. King, K. Rahn, and S. Read. 1997. Use of the flagellar H7 gene as a target in multiplex PCR assays and improved specificity in identification of enterohemorrhagic Escherichia coli s trains. J. Clin. Microbiol. 35: 656662. Geden, C. J. 2006. Visual targets for capture and management of house flies, Musca domestica L. J. Vector Ecol. 31: 152157. Geden, C. J., J. A. Hogsette, and R. D. Jacobs. 1999. Effect of airflow on house fl y (Diptera: Muscidae) distribution in poultry houses. J. Econ. Entomol. 92: 41620. Geden, C. J., D. C. Steinkraus, D. A. Rutz. 1993. Evaluation of two methods for release of Entomophthora muscae (Entomophthorales: Entomophthoraceae) to infect house flies (Diptera: Muscidae) on dairy farms. Environ. Entomol. 20: 12011208. [Genvault] GenVault Corp. 2010. DNA Quantitation: methods and recommendations in use at GenVault. ( www.genvault.com ) Gillespie, J. R. 2002. M odern livestock and poultry production, 6th ed. Thomas Delmare Learning, Albany, NY. Google Earth. 2009. Google Earth. ( http://earth.google.com/downloadearth.html) Goulson, D., L. C. Derwent, M. E. Hanley, D. W. Dunn, and S. R. Abolins. 2005. Predicting calyptrate fly populations from the weather, and probably consequences of climate change. J. Appl. Ecol. 42: 795804. Graham Smith G. S. 1912. An investigation into the possibility of pathoge nic microorganisms being taken up by the larva and subsequently distributed by the fly, pp. 330335. In 41st Ann. Rep. Local Govt. Bd. Suppl. Re p. Med. Off. 19111912. App. B Graham Smith. 1939. .Further observations on the relation of the decline in the number of horse drawn vehicles to the fall in the summer diarrhoea death rate. J. Hyg.39:558562. Gratz, N. G. 1999. Emerging and resurging vector borne diseases. Annu. Rev. Entomol. 44: 5175. Greenberg, B. 1959a. House fly nutrition. II. Comparative s urvival values of sucrose and water. Ann. Entomol. Soc. Am. 53: 125128.

PAGE 194

194 Greenberg, B. 1959b. Persistence of bacteria in the developmental stages of the housefly. I. Survival of enteric pathogens in the normal and aseptically reared host. Am. J. Trop. Med. Hyg. 8: 405411. Greenberg, B. 1959c. Persistence of bacteria in the developmental stages of the housefly. II. Quantitative study of the host contaminant relationship in flies breeding under natural conditions. Am. J. Trop. Med. Hyg. 8: 412416. Green berg, B. 1959d. Persistence of bacteria in the developmental stages of the housefly. III. Quantitative distribution in prepupae and pupae. Am. J. Trop. Med. Hyg. 8: 613617. Greenberg, B. 1959e. Persistence of bacteria in the developmental stages of the h ousefly. IV. Infectivity of the newly emerged adult. Am. J. Trop. Med. Hyg. 8: 61822. Greenberg, B. 1965. Flies and disease. Sci. Am. 213: 9299. Greenberg, B. 1971. Flies and disease, Vol. I ecology, classification and biotic associations. Princeton University Press, Princeton, NJ. Greenberg, B. 1973. Flies and disease, Vol. II biology and disease transmission. Princeton University Press, Princeton, NJ. Greenberg, B., J. A. Kowalski, and M. J. Klowden. 1970. Factors affecting the transmission of Salmonella by flies: natural resistance to colonization and bacterial interference. Infect. Immun. 2: 800809. Greenquist, M. A., J. S. Drouillard, J. M. Sargeant, B. E. Depenbusch, X. Shi, K. F. Lechtenberg, and T. G. Nagaraja. 2005. Comparison of rectoa nal mucosal swab cultures and fecal cultures for determining prevalence of Escherichia coli O157:H7 in feedlot cattle. Appl. Environ. Microbiol. 71: 6431 6433. Grif K., M. P. Dierich H. Karch, F. Allerberger. 1998. Strainspecific differences in the amount of Shiga toxin released from enterohaemorrhagic Escherichia coli O157 following exposure to subinhibitory concentrations of antimicrobial agents. Eur J Clin Microbiol. Infect Dis 17: 761766. Grbel, P., J. S. Hoffman, F. K. Chong, N. A. Burstein, C. Mepani, and D. R. Cave. 1997. Vector potential of houseflies ( Musca domestica ) for Helicobacter pylori J. Clin. Microbiol. 35: 13001303. Gullan, P. J. and P. S. Cranston. 2000. The insects: an outline of entomology, 2nd ed. Blackwell Science, Osney Mead, Oxford. Hagler, J. R. and C. G. Jackson. 2001. Methods for marking insects: current techniques and future prospects. Annu. Rev. Entomol. 46: 511543.

PAGE 195

195 Hald, B., H. Skovgard, D. D. Bang, K. Pedersen, J. Dybdahl, J. B. Jespersen, and M. Madsen. 2004. Flies and Campylobacter infection of broiler flocks. Emerg. Infect. Dis. 10: 14901492. Halverson, M. 2000. The price we pay for corporate hogs. Institute for Agriculture and Trade Policy (IATP) Report. July 2000, 248 pp. ( http://www.iatp.org/ hogreport ) Hammer O. 1941. Biological and ecological investigations on flies associated with pasturing cattle and their excrement. Vidensk. Meddr. Dansk. Naturh. Foren. 105: 5257. Hancock, D. D., T. E. Besser, M. L. K insel, P. I. Tarr, D. H. Rice, and M. G. Paros. 1994. The prevalence of Escherichia coli O 157:H7 in dairy and beef cattle in Washington State. Epidemiol. Infect. 113: 199207. Hancock, D., D. Rice, L. Thomas, D. Dargataz, and T. Besser. 1997. Epidemiology of Escherichia coli O157 in feedlot cattle. J. Food Prot 60: 462465. Hanec, W. 1956. A study of the environmental factors affecting dispersion of house flies ( Musca domestica L.) in a dairy community near Fort Whyte, Manitoba. Can. Entomol. 88: 270272. Hatch Jr., E. 1911. The housefly as a carrier of disease. Ann. Am. Acad. Political Soc. Sci. 37: 168179. Haupt, A. and J. R. Busvine. 1968. The effect of overcrowding on the size of houseflies ( Musca domestica L.). Trans. R. Entomol. Soc. Lond. 120: 297311. Hawley, J. E., L. R. Penner, S. E. Wedberg, and W. Kulp. 1951. The role of the house fly, ii, in the multiplication of certain enteric bacteria. Am. J. Trop. Med. Hyg. 31: 572582. Heuvelink, A. E. 2003. Review of media for the isolation of diarr hoeagenic Escherichia coli pp. 229247. In J. E. L. Corr y, G. D. W. R. Curtis, and M. E. Baird [eds.]. Handbook of culture media for food microbiology. Elsevier Science, Amsterdam. Hewitt, C. G. 1 914. The house fly, its structure, habits, development, re lation to disease control. University Press, Cambridge, UK. Hibbing, M. E., C. Fuqua, M. R. Parsek, and S. Brook Peterson. Bacterial competition: surviving and thriving in the microbial jungle. Nature Rev. Microbiol. 8: 1525. Hoffmann, J. A. and C. Hetr u. 1992. Insect defensins: inducible antibacterial peptides. Immunol. Today 13: 411415. Hoffman, R. A. and A. W. Lindquist. 1951. Studies on treatment of flies with radioactive phosphorus. J. Econ. Entomol. 44: 471473.

PAGE 196

196 Hogsette, J. A. 1983. An attractant self marking devise for marking field populations of stable flies with fluorescent dusts. J. Econ. Entomol. 76: 510514. Hogsette, J. A. 1984. Effect of flourescent dust color on the attractiveness of attractant self marking devises to the stable fly ( Diptera: Muscidae). J. Econ. Entomol. 77: 130132. Hogsette, J. A. 1992. New diets for production of house flies and stable flies (Diptera: Muscidae) in the laboratory. J. Econ. Entomol. 85: 22912294. Hogsette, J. A. 1996. Development of house flies (Di ptera:Muscidae) in sand containing varying amounts of manure solids and moisture. J. Econ. Entomol. 89: 940945. Hogsette, J. A. 2008. Ultraviolet light traps: design affects attraction and capture. In Proc. 6th Int. Conf. Urb. Pests, Hungary, 4 pp. OOK P ress Kft., Hungary. Hogsette, J. A., D. A. Carlson, and A. S. Nejame. 2002. Development of granular boric acid sugar baits for house flies (Diptera: Muscidae). J. Econ. Entomol. 95: 11101112. Hogsette, J. A., R. Farkas, and C. Thurczy. 2001. Hymenopter an pupal parasitoids recovered from house fly and stable fly (Diptera: Muscidae) pupae collected on livestock facilities in Southern and Eastern Hungary. Environ. Entomol. 30: 107111. Hogsette, J. A., and R. D. Jacobs. 1999. Failure of Hydrotaea aenescen s a larval predator of the house fly, Musca domestica L., to establish in wet poultry manure on a commercial farm in Florida, USA. Med. Vet. Entomol. 13: 349354. Hogsette, J. A., R. D. Jacobs, and R. W. Miller. 1993. The sticky card: device for studying the distribution of adult house fly (Diptera: Muscidae) populations in closed poultry houses. J. Econ. Entomol. 86: 450454. Hogsette, J. A. and J. P. Ruff. 1990. Comparative attraction of four different fiberglass traps to various age and sex classes of stable fly (Diptera: Muscidae) adults. J. Econ. Entomol. 83: 883886. Hogsette, J. A. and F. Washington. 1995. Quantitative mass production of Hydrotaea aenescens (Diptera: Muscidae). J. Econ. Entomol. 88: 12381242. Holland, J. L., L. Louie, A. E. Si mor, and M. Louie. 2000. PCR detection of Escherichia coli O157:H7 directly from stools: Evaluation of commercial extraction methods for purifying fecal DNA. J. Clin. Microbiol. 38: 41084113. Holzapfel, E. P. and J. C. Harrell. 1968. Transoceanic disper sal studies of insects. Pacific Insects 10: 115153.

PAGE 197

197 Howard, L. O. 1900. A contribution to the study of the insect fauna of human excrement (with especial reference to the spread of typhoid fever by flies). Proc. Wash. Acad. Sci. 2: 541604. Howard, L. O. 1910. The house fly, disease carrier: an account of its dangerous activities and of the means of destroying it. Frederick A. Stokes Co. Publ. New York, NY. Hsu, C., T. Tsai, and T. Pan. 2005. Use of the duplex TaqMan PCR system for detection of Shiga l ike toxinproducing Escherichia coli O157. J. Clin. Microbiol. 43: 26682673. Hu, Y., Q. Zhang, and J. C. Meitzler. 1999. Rapid and sensitive detection of Escherichia coli O157:H7 in bovine faeces by a multiplex PCR. J. Appl. Microbiol. 87: 867876. Huss ein, H. S. and T. Sakuma. 2005. Prevalence of Shiga toxinproducing Escherichia coli in dairy cattle and their products. J. Dairy Sci. 88: 450466. Hussein, H. S., B. H. Thran, M. R. Hall, and W. G. Kvasnicka. 2003. Verotoxinproducing Escherichia coli in culled beef cows grazing rangeland forages Exp. Biol. Med. 228: 352357. Hutchison R. H. 1916. Notes on the preoviposition of the house fly, Musca domestica L. U. S. Dept. Agr. Bull., Washington, D. C. 345: 116. [ I OWH] Institute for One World Health. 2008. Diarrheal diseases fact sheet, ( http://www.oneworldhealth.org/diseases/diarrhea.php) Islam, M., M. P. Doyle, S. C. Phatak P. Millner, and X. Jiang. 2004. Persistence of enteroh emorrhagic Escherichia coli O157:H7 in soil and on leaf lettuce and parsley grown in fields treated with contaminated manure composts or irrigation water. J. Food Prot. 67: 13651370. Islam, M., M. P. Doyle, S. C. Phatak P. Millner, and X. Jiang. 2005. Survival of Escherichia coli O157:H7 in soil and on carrots and onions grown in fields treated with contaminated manure composts or irrigation water. Food Microbiol. 22: 63 70. [ ITIS ] Integrated Taxonomic Information System. 2008. Musca domestica Linnaeus 1758. ITIS Report. ( http://www.itis.gov ) Janda, J. M. and S. L. Abbott. 2006. The Enterobacteria, 2nd edition. ASM Press, Washington, D. C. Jiang, X., J. Morgan and M. P. Doyle 2002.Fate of Escherichia coli O157: H7 in manure amended soil. Appl. Environ. Microbiol. 68: 2605 2609. Johnson, C. G. 1966. A functional system of adaptive dispersal of flight. Annu. Rev. Entomol. 11: 233260.

PAGE 198

198 Johnson, C. G. 1969. Migration and dispersal of insects by flight. Methuen and Co., London, UK. Johnson, J. R. 2002. Evolution of pathogenic Escherichia coli pp. 5577. In M. S. Donnenberg [ed.], Escherichia coli : Virulence mechanisms of a versatile pathogen. Academic Press, San Diego, CA. Johnson, R. P., J. B. Wilson, P. Michel, K. Rahn, S. A. Renwick, C. L. Gyles, and J. S. Spika. 1999. Escherichia coli O157 in farm animals. CABI Publishing, Wallingford, Oxon, UK. Jones, C. J., J. A. Hogsette, R. S. Patterson, D. E. Milne, G. D. Propp, J. F. Milio,, L G. Rickard, and J. P. Ruff. 1991. Origin of stable flies (Diptera: Muscidae) on West Florida beaches: electrophoretic analysis of dispersal. J. Med. Entomol. 28: 787795. Karch, H., C. Janetzke Mittmann, S. Aleksic, and M. Datz. 1996. Isolation of enterohemorrhagic Escherichia coli O157 strains from patients with hemolytic uremic syndrome by using immunomagnetic separation, DNA based methods, and direct culture. J. Clin. Microbiol. 34: 516519. Karmali, M. 1989. Infection by verocytotoxinproducing Escherichia coli Clin. Microbiol Rev. 2: 1538. Kaufman, P. E. 2002. Dairy pest management, arthropods, pp. 181183. In D. Pimentel [Ed.] Encyclopedia of pest m anagement Vol. 1. Marcel Dekker, Inc. Kaufman, P. E., M. Burgess and D. A. Rutz. 2002. Population dynamics of manure inhabi ting arthropods under an integrated pest management (IPM) program in New York poultry program in New York poultry facilities 3 case studies. J. Appl. Poul. Sci. Res. 11: 90103. Kaufman, P. E., S. J. Long, and D. A. Rutz. 2001a. Impact of exposure lengt h and pupal source on Muscidifurax raptorellus and Nasonia vitripennis (Hymenoptera: Pteromalidae) parasitism in a New York poultry facility. J. Econ. Entomol. 94: 9981003. Kaufman, P. E., S. J. Long, D. A. Rutz, and J. K. Waldron. 2001b. Parasitism rates of Muscidifurax raptorellus and Nasonia vitripennis (Hymenoptera: Ptermolidae) after individual and paried releases in New York poultry facilities. J. Econ. Entomol. 94: 593598. Kaufman, P. E., C. Reasor, D. A. Rutz, J. K. Ketzis, and J. J. Ahrends. 2005a. Evaluation of Beauveria bassiana applications against adult house flies, Musca domestica in commercial caged layer poultry facilities in New York state. Biol. Control 33: 360 367. Kaufman, P. E., D. A. Rutz, and S. Frisch. 2005b Large sticky traps for capturing house flies and stable flies in dairy calf greenhouse facilities. J. Dairy Sci. 88: 176 181.

PAGE 199

199 Kaufman, P. E., J. G. Scott, and D. A. Rutz. 2001. Monitoring insecticide resistance in house flies (Diptera: Muscidae) from New York dairies. Pest M anag. Sci. 57: 514521. Keen, J. E., T. F.Wittum, J. R. Dunn, J. L. Bono, and L. M. Durso. 2006. Shiga toxigenic Escherichia coli O157 in agricultural fair livestock, United States. Emerg. Infect. Dis. 12: 780786. Klein, E. J., J. R. Stapp, M. A. Neill, J. M. Besser, M. T. Osterholm, and P. I. Tarr. 2004. Shiga toxin antigen detection should not replace Sorbitol MacConkey agar screening of stool specimens. J. Clin. Microbiol. 42: 44164417. Kobayashi, M., T. Sasaki, N. Saito, K. Tamur, K. Suzuki, H. Wat anabe, and N. Agui 1999. Houseflies: not simple mechanical vectors of enterohemorrhagic Escherichia coli O157:H7. Am. J. Trop. Med. Hyg. 61: 625629. Kosek, M., C. Bern, and R. L. Guerrant. 2003. The global burden of diarrhoeal disease, as estimated fr om studies published between 1992 and 2000. Bull. World Health Organ. 81: 197204. Kovacs Sr., F. I. Medveczky, L. Papp and E. Gondar. 1990. Role of prestomal teeth in feeding of the house fly, Musca domestica (Diptera; Muscidae). Med. Vet. Entomol. 4: 331335. Krafsur, E. S., M. A. Cummings, M. A. Endsley, J. G. Marquez, and J. D. Nason. 2005. Geographic differentiation in the house fly estimated by microsatellite and mitochondria variation. J. Hered. 96: 502512. Kristiansen, K. and O. Slovmand. 1985. A method for the study of population size and survival rate of houseflies. Entomol. Exp. Appl. 38: 145 150. Kr ger, A., P. M. A. Lucchesi, and A. E. Parma. 2007. Evaluation of vt 2subtyping methods for identifying vt 2g in verotoxigenic Escherichia coli J. Med. Microbiol. 56: 14741478. Kudva, I. T., K. Blanch, and C. J. Hovde. 1998. Analysis of Escherichia coli O157:H7 survival in ovine or bovine manure and manure slurry. Appl. Environ. Microbiol. 64: 31663174. Kumar, M. and G. G. Carmichael. 1998. Antisense RNA: Function and fate of duplex RNA in cells of higher eukaryotes. Microbiol. Mol. Biol. Rev. 62: 14151434. Lahti, E., O. Ruoho, L. Rantala, M. Hanninen, and T.Honkanen Buzalski. 2003. Longitudinal study of Escherichia coli O157 in a cattle f inishing unit. Appl. Environ. Microbiol. 69: 554561.

PAGE 200

200 Lazarus, W., D. A. Rutz, R. W. Miller, and D. A. Brown. 1989. Costs of existing and recommended manure management practices for house fly and stable fly (Diptera: Muscidae) control on dairy farms. J. Econ. Entomol. 82: 11451151. Ledingham, J. C. G. 1911. On the survival of specific microorganisms in pupae and imagines of Musca domestica raised from experimentally infected larvae. Experiments with B. typhosus J. Hyg. 11: 333340. LeJeune, J. T., D. D. Hancock, and T. E. Besser. 2006. Sensitivity of Escherichia coli O157 detection in bovine feces assessed by broth enrichment followed by immunomagnetic separation and direct plating methodologies. J. Clin. Microbiol. 44: 872875. Levin, S. A and V. A ndreasen 1999. Disease transmission dynamics and the evolution of antibiotic resistance in hospitals and communal set tings. Proc. Natl. Ac a d. Sci. 96: 800 801. Lole, M. J. 2005. Nuisance flies and landfill activities: an investigation at a West Midlands landfill site. Waste Manag. Res. 23: 420428. Lysyk, T. J. 1993. Seasonal abundance of stable flies and house flies (Diptera: Muscidae) in dairies in Alberta, Canada. J. Med. Entomol. 30: 888895. Lysyk, T. J. and R. C. Axtell. 1985. Comparison of baited jug trap and spot cards for sampling house fly, Musca domestica (Diptera: Muscidae), populations in poultry houses. Environ. Entomol. 14: 815819. Lysyk, T. J. and R. C. Axtell. 1986. Field evaluation of three methods for monitoring populations of house flies ( Musca domestica ) (Diptera: Muscidae) and other filth flies in three types of poultry housing systems. J. Econ. Entomol. 79: 144151. Lysyk, T. J., L. D. Kalischuk Tymensen, L. B. Selinger, R. C., Lancaster, L. Wever, and K. J. Cheng. 1999. Rearing stable fly larvae (Diptera: Muscidae) on an egg yolk medium. J. Med. Entomol. 36: 382388. MacLeod, J. and J. Donnelly. 1960. Natural features and blowfly movement. J. Anim. Ecol. 29: 8593. Macloskie, G. 1880. The proboscis of the house fly. Am. Natur. 14: 153 161. Macovei, L. and L. Zurek. 2007. Influx of enterococci and associated antibiotic resistance and virulence genes from readyto eat food to the human digestive tract. Appl. Environ. Microbiol. 73: 67406747. Macovei L., B. Miles, and L. Zurek. 2008. Potential of houseflies to contaminate readyto eat food with antibiotic resistant enterococci. J. Food Prot. 71: 435439.

PAGE 201

201 Madigan, M. T. and J. M. Martinko. 2006. Brock biology of m icroorganisms. Benjamin Cummings, San Francisco, CA. Mai, V., C. R. Braden, J. Heckendorf, B. Pironis, and J. M. Hirshon. 2006. Monitoring of stool microbiota in subjects with diarrhea indicates distortions in composition. J. Clin. Microbiol. 44: 4550455 2. Majalija, S., H. Segal, F. Ejobi, and B. G. Elisha. 2008. Shi ga toxin gene containing Escherichia coli from cattle and diarrheic children in the pastoral systems of southwestern Uganda. J. Clin. Microbiol. 46: 352 354. Malik, A., N. Singh, and S. Satya. 2007. House fly ( Musca domestica ): A review of control strategies for a challenging pest. J. Environ. Sci. Health, Part B 42: 453469. Manafi, M. 2003. Media for detection and enumeration of 'total' Enterobacteriaceae, coliforms and Escherichia coli from water and foods, Handbook of cul ture media for food microbiology. Elsevier Science, Amsterdam. Matthews, L., J. C. Low, D. L. Gally, M. C. Pearce, D. J. Mellor, J. A. P. Heesterbeek, M. Chase Topping, S. W. Naylor, D. J. Shaw, S. W. J. Reid, G. J. Gunn, and M. E. J. Woolhouse. 2005. Het erogeneous shedding of Escherichia coli O157 in cattle and its implications for control. Proc. Natl. Acad. Sci. 103: 547552. Matthysse, J. G. 1945. Observations on housefly overwintering. J. Econ. Entomol. 38: 493494. McKay, T., C. D. Steelman, S. M. B razil, and A. L. Szalanski. 2007. Sustained mass release of pupal parasitoids (Hymenoptera: Pteromalidae) for control of Hydrotaea aenescens and Musca domestica (Diptera: Muscidae) in broilerbreeder poultry houses in Arkansas. J. Agr. Urban Entomol. 24: 6785. Mead, P. S. and P. M. Griffin. 1998. Escherichia coli O157:H7. Lancet 352: 12071212. Mead, P. S., L. Slutsker, V. Dietz, L. F. McCraig, J. S. Bresee, C. Shapiro, P. M. Griffin, and R. V. Tauxe. 1999. Foodrelated illness and death in the United States. Emerg. Infect. Dis. 5: 607625. [ MEDIC] Medical Education Information Center. 1995. Department of Path ology and Laboratory Medicine's Enterobacteriaceae. ( http://medic.med.uth.tmc.edu/ path/00001500.htm ) Meerburg, B. G., H. M. Vermeer, and A. Kijlstra. 2007. Controlling risks of pathogen transmission by flies on organic pig farms. Outlook Agric. 36: 193197. Mennigmann, H. D. 1979. Storage death at low temperature (18 C) of strains of Escherichi a coli with different repair capacities. J. Gen. Microbiol. 112: 207210.

PAGE 202

202 Miles, E. J. 1959. Disinfestation and control of pests on refuse tips. J. Roy. Soc. Prom. Health 79: 268273. Milio, J., C. S. Lofgren, and D. F. Williams. 1988. Nuptial flight studies of field collected colonies of Solenopsis invicta Buren pp. 419431. In J. C. Trager [ ed.] Advances in Myrmecology, E. J. Brill, Leiden, NY Moon, R. D. 2002. Muscid flies (Muscidae) pp. 279302. In Mullen, G. and L. Durden [eds.].Medical and vet erinary entomology. Elsevier Science Academic Press, London, UK. Moon, R. D. and H. J. Meyer. 1985. Nonbiting flies pp. 65 82. In Williams, R. E., R. D. Hall, A. B. Broce, and P. J. Scholl [ eds. ] Livestock entomology. John Wiley and Sons, New York, NY. Morabito, S., G. DellOmo, U. Agrimi, H. Sc hmidt, H. Karch, T. Cheasty, and A. Caprioli. 2001. Detection and characterization of Shiga toxinproducing Escherichia coli in feral pigeons. Vet. Microbiol. 82: 275283. Morgan, N. O. and L. G. Pickens. 1978. XI. House flies and other nonbiting flies (Family Muscidae). United States Department of Agric ulture Handbook: Surveillance and collection of arthropods of veterinary importance 518: 7276. Morgan, P. B., D. E. Weidhaas, and G. C. LaBrecque. 1979. Host parasite relationship of the house fly, Musca domestica L., and the microhymenopteran pupal parasite, Muscidifurax raptor Girault and Sanders (Diptera: Muscidae and Hymenoptera: Pteromalidae). J. Kansas Entomol. Soc. 52: 276281. Mullen, G. and L. Durden. 2002. Medical and veterinary entomology. Elsevier Science Academic Press, London, UK. Murinda, S. E. and S. P. Oliver. 2006. Physiologic and molecular markers for detection of Shiga toxin producing Escherichia coli serotype O26 strains. Foodborne Path. Di s. 3: 163177. Murvosh, C. M., R. L. Fye, and G. C. Labrecque. 1964. Studies on the mating behavior of the house fly, Musca domestica L. Ohio J. Sci. 4: 264271. Murvosh, C. M. and C. W. Thaggard. 1966. Ecological studies of the house fly. Ann. Entomol. Soc. Am. 59: 533547. Nash, J. T. C. 1909. House flies as carriers of disease. J. Hyg. 9: 141 169. Nash, J. T. C. 1913. Range of flight of Musca domestica Lancet 182:15851586.

PAGE 203

203 Nataro, J. P. and J. B. Kaper. 1998. Diarrheagenic Escherichia coli Clin Microbiol. Rev. 11: 132201. Nation, J. L. 2002. Insect physiology and biochemis try. CRC Press, Boca Raton, FL. Nayduch, D., A. Honko, G. P. Noblet, and F. Stutzenberger. 2001. Detection of Aeromonas caviae in the common housefly Musca domestica by cul ture and polymerase chain reaction. Epidemiol. Infect. 127: 561 566. Nayduch, D., G. P. Noblet, and F. J. Stutzenberger. 2005. Fate of bacteria, Aeromonas caviae in the midgut of the housefly, Musca domestica Invert. Biol. 124: 7478. Naylor, S. W., J. C. Low, T. E. Besser, A. Mah ajan, G. J. Gunn, M. C. Pearce I. J. McKendrick, D. G. E. Smith, and D. L. Gally. 2003. Lymphoid follicle dense mucosa at the terminal rectum is the principal site of colonization of enterohemorrhagic Escherichia coli O157:H7 in the bovine host. Infect. Immun. 71: 15051512. Nazni, W. A., H. Luke, W. M. Wan Rozita, A. G. Abdullah, I. Sa'diyah, A. H. Azahari, I. Zamree, S. B. Tan, H. L. Lee, and M. A. Sofian. 2005. Determination of the flight range and dispersal of the house fl y, Musca domestica (L.) using mark release recapture technique. Trop. Biomed. 22: 5361. [NCBI] National Center for Biotechnology Information 2008. NCBI Taxonomy. ( http://www.chem.missouri.edu/TannerGroup/people/white/NCBI%20taxonomy%20bro wser/ecoli.htm). Neidhardt, F. C., R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbar ger [eds.]. 1996. Escherichia coli and Salmonellae: cellular and molecular biology 2nd edition. ASM Press, Washington, DC. New, T. R. 1998. Invertebrate surveys for conservation. Oxford University Press, New York, NY. Nmorsi, O. P. G., G. Agbozele, and N. C. D. Ukwandu. 2007. Some aspects of epidemiology of filth flies: Musca domestica Musca domestica vicina, Drosophila melanogaster and associated bacteria pathogens in Ekpoma, Nigeria. Vector Borne Zoonotic Dis. 7: 107117. Ogden, I. D., N. F. Hepburn and M. MacRae. 2001. The optimization of isolation media used in immunomagnetic separation methods for the detection of Escherichia coli O157 in foods. J. Appl. Microbiol. 91: 17. Olsen, A. R. 1998. Regulatory action criteria for filth and other extran eous materials: III. Review of flies and foodborne enteric disease. Regul. Toxicol. Pharmacol. 28: 199211.

PAGE 204

204 Olson, T. A. and R. G. Dahms. 1945. Control of housefly breeding in partly digested sewage sludge. J. Econ. Entomol. 38: 602604. Omisakin, R., M. MacRae, I. D. Ogden, and N. J. C. Strachan. 2003. Concentration and prevalence of Escherichia coli O157 in cattle feces at slaughter. Appl. Environ. Microbiol. 69: 24442447. Ooka, T., J. Terajima, M. Kusumoto, A. Iguchi, K. Kurokwa, Y. Ogura, M. Asadulghani, K. Nakayama, K. Murase, M. Ohnishi, S. Iyoda, H. Watanabe, and T. Hayashi. 2009. Development of a multiplex PCR based rapid typing method for enterohemorrhagic Escherichia coli O157 strains. J. Clin. Microbiol. 47: 28882894. Oporto, B., J. I. Esteb an, G. Aduriz R. A. Juste, and A. Hurtado. 2008. Escherichia coli O157:H7 and nonO157 Shiga toxin producing E. coli in healthy cattle, sheep and swine herds in Northern Spain. Zoonotic Publ. Health 55: 7381. Orth, D., K. Grif, M. P. Dierich, and R. Wu rzner. 2007. Variability in tellurite resistance and the ter gene cluster among Shiga toxinproducing Escherichia coli isolated from humans, animals and food. Res. Microbiol. 158: 105111. Osek, J. 2001. Molecular characterisation of Shiga toxin producing Escherichia coli O157 strains isolated in Poland. Int. J. Food Microbiol. 70: 175177. Pao, S., D. Patel, A. Kalantari, J. P. Tritschler, S. Wildeus, and B. L. Sayre. 2005. Detection of Salmonella strains and Escherichia coli O157:H7 in feces of small ruminants and their isolation with various media. Appl. Environ. Microbiol. 71: 21582161. Parker, R. R. 1916. Dispersion of Musca domestica Linnaeus under city conditions in Montana. J. Econ. Entomol. 9: 325354. Parkes, L. C. 1911. The common house fly ( M usca domestica ). J. Roy. Soc. Prom. Health 32: 4. Paton, A. W. and J. C. Paton. 1998. Detection and characterization of Shiga toxigenic Escherichia coli by using multiplex PCR assays for stx1, stx2, eaeA enterohemorrhagic E. coli hlyA rfbO111, and rfbO1 57. J. Clin. Microbiol. 36: 598602. Pearce, M. C., D. Fenlon, J. C. Low, A. W. Smith, H. I. Knight, J. Evans, G. Foster, B. A. Synge, and G. J. Gunn. 2004. Distribution of Escherichia coli O157 in bovine fecal pats and its impact on estimates of the prev alence of fecal shedding. Appl. Environ. Microbiol. 70: 57375743. Pedigo, L. P. and G. D. Buntin [eds.] 1994. Handbook of sampling methods for arthropods in agriculture. CRC Press, Boca Raton, FL Pell, A. N. 1997. Manure and microbes: public and anima l health problems? J. Dairy Sci. 80: 26732681.

PAGE 205

205 Peppler, H. J. 1944. Usefulness of microorganisms in studying dispersal of flies. Bull. U.S. Army Med. Dept. 75: 121122. Perna, N. T., J. D. Glasner, V. Burland and G. Plunkett III. 2002. The genomes of E scherichia coli K 12 and pathogenic E. coli pp. 353. In Donnenberg, M. S. [ed.]. Escherichia coli : Virulence mechanisms of a versatile pathogen. Academic Press, San Diego, CA. Perotti, M. A. and T. J. Lysyk. 2003. Novel growth media for rearing larval h orn flies, Haematobia irritans (Diptera: Muscidae). J. Med. Entomol. 40:2229. Petridis, M., M. Bagdasarian, M. K. Waldor, and E. Walker. 2006. Horizontal transfer of Shiga toxin and antibiotic resistance genes among Escherichia coli strains in house fly (Diptera: Muscidae) gut. J. Med. Entomol. 43: 288295. Pickens, L. G., N. O. Morgan, J. G. Hartsock, and J. W. Smith. 1967. Dispersal patterns and populations of the house fly affected by sanitation and weather in rural Maryland. J. Econ. Entomol. 60: 12501255. Pickens L. G., N. O. Morgan, and R. W. Miller. 1972. Comparison of traps and other methods for surveying density of populations of house flies in dairy barns. J. Econ. Entomol. 65: 144145. Pickens, L. G. 1989. Factors affecting the distance of scatter of house flies (Diptera: Muscidae) from electrocuting traps. J. Econ. Entomol. 82: 149151. Pimentel, D. W. P. Nagel, and J. L. Madden. 1963. Spacetime structure of the environment and the survival of parasite host systems. Am. Nat. 97: 141 167. P ollack, K. 2005. Enhanced surveillance of haemolytic uraemic syndrome and other thrombotic microangiopathies in Scotland, 2003 2004. Euro Surveill. 10:pii=2708. ( http://www.eurosurveillance.org/ViewArticle.aspx?ArticleId=2708) Porter, J., K. Mobbs, C. A. Hart, J. R. Saunders, R. W. Pickup, and C. Edwards. 1997. Detection, distribution and probable fate of Escherichia coli O157 from asymptomatic cattle on a dairy farm. J. Appl Microbiol. 83: 297 306. Powell, D. 2008. The human face of E. coli O157:H7: 3year old died in 2000. ( http://barfblog.foodsafety.ksu.edu/blog/138170/08/06/15/humanfacee coli O157H7 3year old died 2000) Quarterman, K. D., W. Mathis, and J. W. Kilpatrick. 1954. Urban fly dispersal in the area of Savannah, Georgia. J. Econ. Entomol. 47: 405412. Rasmussen, M. A., T. L. Wickman, W. C. Cray Jr., and T. A. Casey. 1999. Escherichia coli O157:H7 and the rumen environment, pp. 3949. In S tewart, C. S., and H. J. Flint [ eds. ] Escherichia coli O157 in farm anim als. CABI, New York, NY.

PAGE 206

206 Reisen, W. K. 2010. Landscape epidemiology of vector borne diseases. Annu. Rev. Entomol. 55: 461483. Renter, D. G., S. L. Checkley, J. Campbell, and R. King. 2004a. Shiga toxin producing Escherichia coli in the feces of Alberta feedlot cattle. Can. J. Vet. Res. 68: 150153. Renter, D. G., J. M. Sargeant, L. L. Hungerford. 2004b Distribution of Escherichia coli O157:H7 within and among cattle operations in pasture based agricultural areas. Am. J. Vet. Res. 65: 13671376. Rice, D. H., K. M. McMenamin, L. C. Pritchett, D. D. Hancock, and T. E. Besser. 1999. Gene tic subtyping of Escherichia coli O157 isolates from 41 Pacific Northwest USA cattle farms. Epidemiol. Infect. 122: 479484. Rice, D. H., H. Q. Sheng, S. A. Wynia, and C. J. Hovde. 2003. Rectoanal mucosal swab culture is more sensitive than fecal culture and distinguishes Escherichia coli O157:H7colonized cattle and those transiently shedding the same organism. J. Clin. Microbiol. 41: 49244929. Riemann, J. G., D. J. Moen, and B. J. Thorson. 1967. Female monogamy and its control in houseflies. Insect Phy siol. 13: 407408. Riley L.W., R. S. Remis, S. D. Helgerson, H. B. McGee, J. G. Wells, B. R. Davis R. J. Hebert, E. S. Olcott, L. M. Johnson, N. T. Hargrett, P. A. Blake, and M. L. Cohen 1983. Hemorrhagic colitis associated with a rare Escherichia coli serotype. N. Engl. J. Med. 308: 681685. Rodriguez, J. G. 1966. Axenic Arthropoda: current status of research and future possibilities. Ann. New York Acad. Sci. 139: 53 64. Rohde, M. 2008. Deal reached in E. coli death. Milwaukee Wisconsin Journal Sentinal Online. ( http://www.jsonline.com/news/milwaukee/29482674.html Rosef, O. and G. Kapperud. 1983. House flies ( Musca domestica ) as possible vectors of Campylobacter fetus subsp. jejuni. Appl. Environ. Microbiol. 45: 381383. Ruiu, L., A. Satta, and I. Floris. 2007. Susceptibility of the house fly pupal parasitoid Muscidifurax raptor (Hymenoptera: Pteromalidae) to the entomopathogenic bacteria Bacillus thuringiensis and Brevibacillus late rosporus. Biol. Control 43: 188194. Rutz, D. A., C. J. Geden, C. W. Pitts. 1994. Pest management recommendations for dairy cattle. Cornell University and Penn. State Coop. Ext., Ith a ca, NY. Sacc, G. 1958. Research on speciation in the housefly. VI. Nat ural hybridism & experimental hybridism between subspecies of Musca domestica L. Rend. Ist. Sup. Sanit. 21: 1170 1184.

PAGE 207

207 Sacc, G. 1964. Comparative bionomics in the genus Musca Annu. Rev. Entomol. 9: 341358. Sacc, G. and L. Rivosecchi. 1958. Research on speciation in the housefly. V. Geographic distribution of the subspecies of Musca domestica L. (Diptera: Muscidae). Rend. Ist. Sup. Sanit. 21: 11491189. Sanderson, M.W., J. M. Gay, D. D. Hancock, C. C. Gay, L. K. Fox, and T. E. Besser. 1995. Sensitivity of bacteriologic culture for detection of Escherichia coli O157:H7 in bovine feces. J. Clin. Microbiol. 33: 26162619. Sanderson, M., J. M. Sargeant, X. Shi, T. G. Nagaraja, L. Zurek, and M. J. Alam. 2006. Longitudinal emergence and distribution of E scherichia coli O157 genotypes in a beef feedlot. App. Environ. Microbiol. 72: 76147619. Sapers, G. M. and M. P. Doyle. 2009. Scope of the produce contamination problem., pp. 319. In Solomon, E. B., and K. R. Matthews [ eds. ], The produce contamination probl em: causes and solutions. Academic Press, Elsevier Inc., Burlington, MA Sargeant, J. M., M. W. Sanderson, R. A. Smith, and D. D. Griffin. 2003. Escherichia coli O157 in feedlot cattle feces and water in four major feeder cattle states in the USA. Prev. V et. Med. 61: 127135. Sargeant, J. M., M. W. Sanderson, D. D. Griffin, and R. A. Smith. 2005. Factors associated with the presence of Escherichia coli O157 in feedlot cattle water and feed in the Midwestern USA. Prev. Vet. Med. 66: 207237. Sasaki, T., M Kobayashi, and N. Agui. 2000. Epidemiological potential of excretion and regurgitation by Musca domestica (Diptera: Muscidae) in the dissemination of Escherichia coli O157: H7 to food. J. Med. Entomol. 37: 945949. SAS Institute. 2002. SAS/STAT Users m anual, Ver. 9.1. SAS Institute, Cary, NC. Scaife, H. R., D. Cowan, J. Finney, S. F. Kinghorn Perry, and B. Crook. 2006. Wild rabbits ( Oryctolagus cuniculus ) as potential carriers of verocytotoxin producing Escherichia coli. Vet. Rec. 159: 175178. Schmidtmann, E. T., and P. A. W. Martin. 1992. Relationship between selected bacteria and the growth of immature house flies, Musca domestica, in an axenic test system. J. Med. Entomol. 29: 223235. Schoof, H. F. 1959. How far do flies fly, and what effect does flight pattern have on their control: Pest Control 27: 1618, 20, 22, 66. Schoof, H. F., G. A. Mail, and E. P. Savage. 1954. Fly production sources in urban communities. J. Econ. Entomol. 47: 245254.

PAGE 208

208 Schoof, H. F. and R. F. Siverly. 1954a. Multiple rel ease studies on the dispersion of Musca domestica at Phoenix, Arizona. J. Econ. Entomol. 47: 830838. Schoof, H. F. and R. F. Siverly. 1954b. Urban fly dispersion studies with special reference to movement pattern of Musca domestica Am. J. Trop. Med. Hyg. 3: 539547. Schurrer, J. A., S. A. Dee, R. D. Moon, J. Deen, and C. Pijoan. 2006. Evaluation of three strategies for insect control on a commercial swine farm. J. Swine Health Prod. 14: 76 81. Scudder, H. L. 1947. A new technique for sampling the densi ty of house fly ( Musca domestica ) populations. Publ. Health Rep. 62: 609623. Scudder, H. L. 1949. Some principles of fly control for the sanitarian. Am. J. Trop. Med. Hyg. 29: 609623. Sezonov, G., D. Joseleau Petit, and R. DAri. 2007. Escherichia coli physiology in Luria Bertani broth. J. Bacteriol. 189: 87468749. Sheppard, C. 1983. House fly and lesser fly control utilizing the black soldier fly in manure management systems for caged laying hens. Environ. Entomol. 12: 14391442. Shere, J. A., K. J. Bartlett, and C. W. Kaspar. 1998. Longitudinal study of Escherichia coli O157:H7 dissemination on four dairy farms in Wisconsin. Appl. Environ. Microbiol. 64: 13901399. Shura Bura, B. L., A. D. Shaykov, YE. V. Ivanova, A. YA. Glazunova, M. S. Mitryukova, and K. G. Fedorova. 1956. The migration of synanthropic flies to a town from the open. Meditc. parzitolog. i parazit. boleznc. 4: 368372. Shura Bura, B. L., O. I. Sukhomlinova, and B. I. Isarova. 1962. Use of radioactive tracers as an aid to studying t he ability of synanthropic flies to fly over water obstacles. Ft. Belvoir: Skoda, S. R., G. D. Thomas, and J. B. Campbell. 1993. Abundance of immature stages of the house fly (Diptera: Muscidae) from five areas in beef cattle feedlot pens. J. Econ. Entom ol. 86: 455461. Slutsker, L., A. A. Ries, K. Maloney, J. G. Wells, K. D. Greene, and P. M. Griffin. 1998. A nationwide casecontrol study of Escherichia coli O157:H7 infection in the United States. J. Infect. Dis. 177:962966. Sokal, R. R. and R. L. Sul livan. 1963. Competition between mutant and wildtype housefly strains at varying densities. Ecol. 44: 314322. Southwood, T. R. E. 1966. Ecological methods. Methuen and Co., London, UK.

PAGE 209

209 Stafford III, K. C. 2008. Fly management handbook: A guide to biology, dispersal, and management of the house fly and related flies for farmers, municipalities, and public health officials. The Conn. Ag. Exp. Sta, New Haven, CT., Bull. 1013, May 2008. Stein, W. 1986. Dispersal of insects of public health importance, pp. 242252. In W. Danthanarayana [ed.] Insect Flight: Dispersal and Migration. Springer Verlag, Berlin, Heidelberg. Steinhaus, E. A. 1940. The microbiology of insects: with special reference to the biologic relationships between bacteria and insects. Bact. Rev. 4:1757. Steinhaus, E. A. 1946. Insect microbiology. New York, NY, Hafner Publishing Co. Steinhoff, U. 2005. Who controls the crowd? New findings and old questions about the intestinal microflora. Immunol. Lett. 99:12 16. Steinkraus, D. C., C. J. G eden, and D. A. Rutz. 1993. Prevalence of Entomophthora muscae (Cohn) Fresenius (Zygomycetes: Entomophthoraceae) in house flies (Diptera: Muscidae) on dairy farms in New York, and induction of epizootics. Biol. Control. 5: 405411. Stoetzel, M. B. 1989. Common names of insects and related organisms. Entomol. Soc. of America, College Park, MD. Suda, K. J., B. L. Love, T. J. Gladney, and K. W. Garey. 2003. Health and economic outcomes of hospitalized patients with Clostridium difficile associated diarrhea. Abstr. Intersci. Conf. Antimicrob. Agents Chemother. (43rd meeting, abstract no. K 734). Sukontason, K., K. L. Sukontason, R. C. Vogtsberger, N. Boonchu, T. Chaiwong, and S. Piangjai. 2003. Prestomal teeth of some flies of medical importance. Micron 34: 449452. Sulaiman, S. M. Z. Othman, and A. H. Aziz. 2000. Isolations of enteric pathogens from synanthropic flies trapped in downtown Kuala Lumpar J. Vec. Ecol. 25: 9093. Swadener, C. 1994. Bacillus thuringiensis ( B.t. ). J. Pest. Reform 14: Fall 1994. http://www.safe2use.com/poisons pesticides/pesticides/BtK/btk.htm Szalanski, A. L., C. B. Owens, T. McKay, and C. D. Steelman. 2004. Detection of Campylobacter and Escherichia coli O157:H7 from filth flies by polymerase chain reaction. Med. Vet. Entomol. 18: 241246. Taft, H. M. and H. R. Agee. 1962. A marking and recovery method for use in boll weevil movement studies. J. Econ. Entomol. 55: 10181019. Tarr, P. I. and M. A. Neill. 2001. Escherichia coli O157:H7. Gastroenterol. Clin. Am. 30: 735751.

PAGE 210

210 Taylor, C. E. and R. R. Sokal. 1976. Oscillations in housefly population sizes due to time lags. Ecol. 57: 10601067. Taylor, L. R. 1963. Analysis of the effect of temperature on insects in flight. J. Anim. Ecol. 32: 99117. Teplitski, M., J. D. Barak, and K. R. Schneider. 2009. Human enteric pathogens in produce: unanswered ecological questions with direct implications for food safety. Curr. Op. Biotechnol. 20: 166 171. Te sch, M. J. and W. G. Goodman. 1995. Dissemination of microbial contaminants from house flies electrocuted by five insect light traps. Inter. J. Environ. Health Res. 5: 303309. Thimijan, R. W., L. G. Pickens, N. O. Morgan, and R. W. Miller. 1972. House fl y capture as a function of number of traps in a dairy barn. J. Econ. Entomol. 65: 876877. Thomas, G. D.1993. The influence of beef cattle feedlots on the urban fly problem pp. 116. In Thomas, G. D. and S. R. Skoda [ eds. ] Rural flies in the urban envi ronment? North Central Regional Res. Bull. 335, Institute of Agriculture and Natural Resources Res. Bull. 317. University of Nebraska Agricultural Research Division, Lincoln, NE. Thomas, G. D. and S. R. Skoda. [eds.]. 1993. Rural flies in the urban environment? Research Bulletin 317. North Central Regional Research Publication No. 335. Thompson, F. C. 2009. Nearctic Diptera: twenty years later, pp. 3 46. In T. Pape, D. Bickel, and R. Meier [eds.]. Diptera diversity: status, challenges and tools. Koninklij ke Brill, Leiden, The Netherlands. Thorpe, C. M., J. M. Ritchie, and D. W. K. Acheson. 2002. Enterohemorrhagic and other Shiga toxin producing Escherichia coli pp. 119154. In Donnenberg, M. S. [ed.]. Escherichia coli : Virulence mechanisms of a versatile pathogen. Academic Press, San Diego CA. Tobin, P. C. and C. W. Pitts. 1999. Dispersal of Muscidifurax raptorellus Kogan and Legner (Hymenoptera: Pteromalidae) in a high rise poultry facility. Biol. Control 16: 6872. Todar, K. 2008. Pathogenic E. coli In Online textbook of bacteriology. Univ. WisconsinMadison Dept. Bacteriol. ( http://www.textbookofbacteriology.net/e.coli.html). Tsuda, Y., H. Toshihiko, Y. Higa, K. Hoshino, S. Kasai, T Tomita, H. Kurahashi, and M. Kobayashi. 2009. Dispersal of a blow fly, Calliphora nigribarbis in relation to the dissemination of highly pathogenic avian influenza virus. Jpn. J. Infect. Dis. 62: 294297. Turchin, P. 1998. Quantitative analysis of move ment. Sinauer Associates, Inc. Publ. Sunderland, MA.

PAGE 211

211 Turchin, P., F. J. Odendaal, and M. D. Rausher. 1991. Quantifying insect movement in the field. Environ. Entomol. 20: 955963. Turner Jr., E. C. and P. L. Ruszler. 1989. A quick and simple quantitative method to monitor house fly populations in caged layer houses. Poult. Sci. 68: 833 835. Tutenel, A. V., D. Pierard, D. Vandekerchove, J. Van Hoof, and L. De Zutter. 2003. Sensitivity of methods for the isolation of Escherichia coli O157 from naturally infected bovine faeces. Vet. Microbiol. 94: 341 346. Tyler, H. L. and E. W. Triplett. 2008. Plants as a habitat for beneficial and/or human pathogenic bacteria. Annu. Rev. Phytopathol. 46: 5371. [ UF EHS ] University of Florida, Environmental Health and Safety. 2008. Biological safety. ( http://www.ehs.ufl.edu/bio/ ). Unc, A., and M. J. Goss. 2006. Culturable Escherichia coli in soil mixed with two types of manure. Soil Sci. Soc. Am. J. 70: 763769. [ USDA:APHIS:VS ] United States Department of Agriculture, Animal and Plant Health Inspection Service, Veterinary Service. 1997. An update: Escherichia coli O157:H7 in humans and cattle. USDA:APHIS:VS Centers for Epidemiology and Animal Health. Fort Collins, CO. ( http://www.aphis.usda.gov/animal_health/emergingissues /downloads/ecoupdat.pdf ) [ USDAFSIS] Food Safety and Inspection Service, Office of Public Health Science. 2008a. Detection, isolation and identification of Escherichia coli O157:H7 from meat products. Microbiol. Lab. Guide (MLG) 5.04. ( www. fsis .usda .gov/PDF/ MLG _5_04.pdf ). [ USDAFSIS] Food Safety and Inspection Service, Office of Public Health Science. 2008b. Procedure for the use of Escherichia coli O157:H7 screening tests. Microbiol. Lab. Guide (MLG) 5A.01. ( www. fsis .usda .gov/PDF/ Mlg _5A_01.pdf ) Van Donkersgoed, J., T. Graham, and V. Gannon. 1999. The prevalence of verotoxins, Escherichia coli O157:H7, and Salmonel la in the feces and rumen of cattle at processing. Can. Vet. J. 40: 332338. Van Donkersgoed, J., D. Hancock, D. Rogan, and A. A. Potter. 2005. Escherichia coli O157:H7 vaccine field trial in 9 feedlots in Albe rta and Saskatchewan. Can. Vet. J. 46: 724728. van Zwet, A. A., J. C. Thijs, A. M. D. KooistraSmid, J. Schirm, and J. A. M. Snijder. 1994. Use of PCR with feces for detection of Helicobacter pylori infections in patients. J. Clin. Microbiol. 32: 13461348. Vidovic, S. and D. R. Korber. 2006. Prev alence of Escherichia coli O157 in Saskatchewan

PAGE 212

212 cattle: Characterization of isolates by using random amplified polymorphic DNA PCR, antibiotic resistance profiles, and pathogenicity determinants. Appl. Environ. Microbiol. 72: 43474355. Visetsripong, A., K. Pattaragulwanit, J. Thaniyavarn, R. Matsuura, A. Kuroda, and O. Sutheinkul. 2007. Detection of Escherichia coli O157:H7 vt and rfbO157 by multiplex polymerase chain reaction. Southeast Asian J. Trop. Med. Public Health 38: 8290. Vold, L., A. Holck, Y. Wasteson, and H. Nissen. 2000. High levels of background flora inhibits growth of Escherichia coli O157:H7 in ground beef. Int. J. Food Microbiol. 56: 219225. Wallace, J. S. and K. Jones 1996. The use of selective and differential agars in the isolation of Escherichia coli O157 from dairy herds. J. Appl. Bacteriol. 81: 663668. Wang, L. and P. R. Reeves. 1998. Organization of Escherichia coli O157 O antigen gene cluster and identification of its specific genes. Infect. Immun. 66: 35453551. Watanabe, Y ., K. Ozasa, J. H. Mermin, P. M. Griffin, K. Masuda, S. Imashuku, and T. Sawada. 1999. Factory outbreak of Escherichia coli O157:H7 infection in Japan. Emerg. Infect. Dis. 5: 424428. Watson, D. W., C. J. Geden, S. J. Long, and D. A. Rutz. 1995. Efficacy of Beuveria bassiana for controlling the house fly and stable fly (Diptera: Muscidae) Biol. Control 5: 405411. Watson, D. W., E. L. Nino, K. Rochon, S. Denning, L. Smith, and J. S. Guy. 2007. Experimental evaluation of Musca domestica (Diptera: Muscidae) as a vector of Newcastle disease virus. J. Med. Entomol. 44: 666 671. Watson, D. W. and J. J. Petersen. 1993. Seasonal activity of Entomophthora muscae ( Zygomycetes : Entomophthorales) in Musca domestica L., (Diptera: Muscidae) with reference to temperature and relative humidity. Biol. Control. 3: 2226. Watson, D. W., P.A.W. Martin, and E. T. Schmidtmann. 1993. Egg yolk and bacteria growth medium for Musca domestica (Diptera: Muscidae). J. Med. Entomol. 30: 820823. Weather Underground. 2009. Daily su mmary history for KFLLAKEC8. ( http://www.wunderground.com/weatherstation/WXDailyHistory.asp?ID=KFLLAKEC8) Wellington, W. G. 1945. Conditions governing the distribution of insects in the free atmosphere. Can. Ent. 77: 715. West, L. 1951. The housefly. its natural history, medical importance and control. Comstock Pub., Ithaca, NY.

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213 Wetzel, A. N. and J. T. LeJeune. 2006. Clonal dissemination of Escherichia coli O157:H7 subtypes among dairy farms in Northeast Ohio. Appl. Environ. Microbiol. 72: 26212626. Wilde, J., J. Eiden, and R. Yolken. 1990. Removal of inhibitory substances from human fecal specimens for detection of group A rotaviruses by reverse transcriptase and polymerase chain reactions. J. Clin. Microbiol. 28: 13001307. Wilkes, A., G. E. Bucher, J. W. M. Cameron, and A. S. West Jr. 1948. Studies on the housefly ( Musca domestica L.) I. The biology and large scale production of laboratory populations. Can J Res. Sec. 26: 2656. Williams, D. F. 1973. Sticky traps for sampling populations of Stomoxys calcitrans J. Econ. Entomol. 66: 12791280. Williams, J. R. P. 1973. Dispersal of 32P labelled adult Fannia canicularis Int. Pest Cont. 15: 2022. Williams D. F., C. S. Lofgren, and R. K. Vander Meer. 1990. Fly pupae as attractant carriers for toxic baits for red imported fire ants (Hymenoptera: Formicidae). J. Econ. Entomol. 83: 6773. Willshaw, G. A., J. Thirlwell, A. P. Jones, S. Parry, R. L. Salmon, an d M. Hickey. 1994. Vero cytotoxinproducing Escherichia coli O157 in beefburgers linked to an outbreak of diarrhoea, haemorrhagic colitis and haemolytic uraemic syndrome in Britain. Lett. Appl. Microbiol. 19: 304307. Wilton, D. P. 1963. Dog excrement as a factor in community fly problems. Proc. Hawaiian Entomol. Soc. 18: 311317. Winpisinger, K. A., A. K. Ferketich, R. L. Berry, and M. L. Moeschberger. 2005. Spread of Musca domestica (Diptera: Muscidae), from two caged layer facilities to neighboring residences in rural Ohio. J. Med. Entomol. 42: 732738. Wong, T. T. Y. and M. L. Cleveland. 1970. Flourescent powder for marking deciduous fruit moths for studies of dispersal. J. Econ. Entomol. 63: 338339. Xavier, B. M. and J. B. Russell. 2006. Bacterial competition between a bacteriocin producing and a bacteriocin negative strain of Streptococcus bovis in batch and continuous culture. FEMS Microbiol. Ecol. 58: 317322. Yates, W. W., A. W. Lindquist and J. S. Butts. 1952. Further studies of dispersion of flies tagged with radioactive phosphoric acid. J. Econ. Entomol. 45: 547548.

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214 Yoruk, R., J. A. Hogsette, R. S. Rolle, and M. R. Marshall. 2003. Apple polyphenol oxidase inhibitor(s) from the common house fly ( Musca domestica L.). J. Food Sci. 68: 19421947. Zarchi, A. A. K. and H. Vatani. 2009. A survey on species and prevalence rate of bacterial agents isolated from cockroaches in three hospitals. Vect or Borne Zoonotic Dis. 9: 197 200. Zarrin, M. Z., B. Vazirianzadeh, S. S. Solary, A. Z. Mahmoudabadi, and M. Rahdar. 2007. Isolation of fungi from housefly ( Musca domestica ) in Ahwaz, Iran. Pak. J. Med. Sci. 23: 917919. Zhao, T., M. P. Doyle, J. Shere, and L. Garber. 1995. Prevalence of enterohemorrhagic Escherichia coli O157:H7 in a survey of dairy her ds. Appl. Environ. Microbiol. 61: 12901293. Zurek, L., C. Schal, and W. Watson. 2000. Diversity and contribution of the intestinal bacterial community to the development of Musca domestica (Diptera: Muscidae) larvae. J. Med. Entomol. 37: 924928. Zurek L., S. S. Denning, C. Schal, and D. W. Watson. 2001. Vector competence of Musca domestica (Diptera: Muscidae) for Yersinia pseudotuberculosis. J. Med. Entomol. 38: 333335.

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215 BIOGRAPHICAL SKETCH Roxanne Burrus is an active duty medical entomologist in the United States Navy. After completing her Ph.D at the University of Florida, she will be reporting to Lima, Peru. Previous duty stations have included Bangor, WA and San Diego, CA. Roxanne's family has a history of military service B oth parents James D. Burrus and Dawn Elaine (Marcet) Burrus, served in the Navy, and her maternal grandfather Henry Marcet, served in the Army. She has one sister, Karen Elizabeth Burrus. Roxanne and her family have lived in many different countries, including the United Stat es, Taiwan, Cyprus, and Spain. Roxanne has two bachelors' degrees ; t he first one in mathematics with a minor in computer science from the University of Southern Mississippi (1988) and the second in biology with a pre medical emphasis from the University of Massachusetts at Amherst (2002) She completed a master's degree in medical and urban entomology at the University of Florida (2004). In spare time, which she had very little of while fulfilling the requirements of this Ph .D during the last three years, Roxanne likes to participate in triathlons and half marathons. She was a member of the UF Triathlon team, the Trigators, during both degree programs at the University of Florida