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Potential of Insecticide-Treated Cords and Sprayable Baits for Control of House Flies (Diptera

Permanent Link: http://ufdc.ufl.edu/UFE0021396/00001

Material Information

Title: Potential of Insecticide-Treated Cords and Sprayable Baits for Control of House Flies (Diptera Muscidae)
Physical Description: 1 online resource (85 p.)
Language: english
Creator: Hertz, Jeffrey C
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2007

Subjects

Subjects / Keywords: bait, control, cords, fipronil, fly, house, imidacloprid, impregnated, indoxacarb, sprayable
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, M.S.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: House flies are often controlled using insecticides when the source of infestation can not be located or remedied by non-chemical methods. Historically, house flies have shown a tremendous potential to develop insecticide resistance and with few classes of insecticides currently registered for house fly control, new products and methods need to be evaluated to prevent future control failures. This research evaluated the potential use of two innovative methods to control house flies: fipronil- and indoxacarb-impregnated cords and a sprayable imidacloprid fly bait. For the insecticide-impregnated cord studies, eight various natural and synthetic cords were evaluated to determine which cords were attractive to house flies. Natural cords were more attractive than synthetic cords; the plant-based manila cord was most attractive and the nylon parachute cord was least attractive. The most attractive cords (manila, cotton, wool, nylon, and polypropylene) were treated with 0.1% fipronil or 0.6% indoxacarb and evaluated in the laboratory to determine their effectiveness. All cords were more effective than the impregnated cotton cord except the fipronil-impregnated nylon cord (LT90) and the indoxacarb-impregnated polypropylene cord. The wool cord was the most effective, LT50 (Fipronil = 12.9 h; Indoxacarb = 32.6 h) and LT90 (Fipronil = 22.4 h; Indoxacarb = 51.5 h). The wool cords were impregnated with 0.1% fipronil and 1.2% indoxacarb and evaluated in a controlled field environment with fresh cords and cords that were aged 4 wk. No significant differences were seen between fly count reductions of either treatment. Both treatments reduced fly counts by > 57% by 24 h and > 87% by 48 h with both fresh and aged cords. A reduction in efficacy was seen with aged cords. The new imidacloprid sprayable fly bait formulation was compared against two commonly used dry scatter baits in the laboratory and against a granular imidacloprid paint-on bait in a controlled field setting. Additionally, the sprayable bait was evaluated for use in impregnated cords. No differences were seen in mortality between the three scatter baits in the laboratory or between the imidacloprid baits in the field cages. Both imidacloprid baits reduced fly counts by > 70% in the field within 24 h, but were not effective after treatments were aged for 2 wk. When various cords were treated with 2.5% of the new bait, the wool cord had higher mortality (74%) compared to the other natural and synthetic cords tested. Knockdown recovery was observed with all bait-treated cords in the laboratory, but was not determined to occur in the field cages. The bait-treated cords reduced fly counts by > 82% with fresh cords and cords aged for 4 wk. Impregnated cords and the new sprayable bait should prove to be valuable tools in established fly management programs in urban, agriculture, and military settings. Fipronil and indoxacarb are not currently registered for house flies, but both appear to be effective insecticides for their control.
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 Jeffrey C Hertz.
Thesis: Thesis (M.S.)--University of Florida, 2007.
Local: Adviser: Koehler, Philip G.

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2007
System ID: UFE0021396:00001

Permanent Link: http://ufdc.ufl.edu/UFE0021396/00001

Material Information

Title: Potential of Insecticide-Treated Cords and Sprayable Baits for Control of House Flies (Diptera Muscidae)
Physical Description: 1 online resource (85 p.)
Language: english
Creator: Hertz, Jeffrey C
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2007

Subjects

Subjects / Keywords: bait, control, cords, fipronil, fly, house, imidacloprid, impregnated, indoxacarb, sprayable
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, M.S.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: House flies are often controlled using insecticides when the source of infestation can not be located or remedied by non-chemical methods. Historically, house flies have shown a tremendous potential to develop insecticide resistance and with few classes of insecticides currently registered for house fly control, new products and methods need to be evaluated to prevent future control failures. This research evaluated the potential use of two innovative methods to control house flies: fipronil- and indoxacarb-impregnated cords and a sprayable imidacloprid fly bait. For the insecticide-impregnated cord studies, eight various natural and synthetic cords were evaluated to determine which cords were attractive to house flies. Natural cords were more attractive than synthetic cords; the plant-based manila cord was most attractive and the nylon parachute cord was least attractive. The most attractive cords (manila, cotton, wool, nylon, and polypropylene) were treated with 0.1% fipronil or 0.6% indoxacarb and evaluated in the laboratory to determine their effectiveness. All cords were more effective than the impregnated cotton cord except the fipronil-impregnated nylon cord (LT90) and the indoxacarb-impregnated polypropylene cord. The wool cord was the most effective, LT50 (Fipronil = 12.9 h; Indoxacarb = 32.6 h) and LT90 (Fipronil = 22.4 h; Indoxacarb = 51.5 h). The wool cords were impregnated with 0.1% fipronil and 1.2% indoxacarb and evaluated in a controlled field environment with fresh cords and cords that were aged 4 wk. No significant differences were seen between fly count reductions of either treatment. Both treatments reduced fly counts by > 57% by 24 h and > 87% by 48 h with both fresh and aged cords. A reduction in efficacy was seen with aged cords. The new imidacloprid sprayable fly bait formulation was compared against two commonly used dry scatter baits in the laboratory and against a granular imidacloprid paint-on bait in a controlled field setting. Additionally, the sprayable bait was evaluated for use in impregnated cords. No differences were seen in mortality between the three scatter baits in the laboratory or between the imidacloprid baits in the field cages. Both imidacloprid baits reduced fly counts by > 70% in the field within 24 h, but were not effective after treatments were aged for 2 wk. When various cords were treated with 2.5% of the new bait, the wool cord had higher mortality (74%) compared to the other natural and synthetic cords tested. Knockdown recovery was observed with all bait-treated cords in the laboratory, but was not determined to occur in the field cages. The bait-treated cords reduced fly counts by > 82% with fresh cords and cords aged for 4 wk. Impregnated cords and the new sprayable bait should prove to be valuable tools in established fly management programs in urban, agriculture, and military settings. Fipronil and indoxacarb are not currently registered for house flies, but both appear to be effective insecticides for their control.
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 Jeffrey C Hertz.
Thesis: Thesis (M.S.)--University of Florida, 2007.
Local: Adviser: Koehler, Philip G.

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2007
System ID: UFE0021396:00001


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POTENTIAL OF INSECTICIDE-TREATED CORDS AND SPRAYABLE BAITS FOR
CONTROL OF HOUSE FLIES (DIPTERA: MUSCIDAE)


















By

JEFFREY CONRAD HERTZ


A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE

UNIVERSITY OF FLORIDA

2007

































2007 Jeffrey Conrad Hertz




























To my wonderful family









ACKNOWLEDGMENTS

I would like to thank the United States Navy for granting me the opportunity to pursue a

Master of Science degree in entomology under the Medical Service Corps In-service

Procurement Program: an experience I certainly didn't anticipate upon enlisting. CAPT (ret.)

Gary Breeden, CAPT Stan Cope, CAPT Mark Beavers, and LCDR Pete Obenauer provided

exceptional support and irreplaceable mentorship throughout this entire endeavor. All of which,

I hope will continue throughout the remainder of my career.

Dr. Phil Koehler has been irreplaceable in his role as my academic advisor. His guidance

and leadership through my entire term at the University of Florida has been exceptional. He has

a knack for teaching that is unlike any other and his emphasis on extension and systematic

problem solving will no doubt benefit me in my future role as a military entomologist.

More has been gained from my committee, Drs. Phil Koehler, Richard Patterson and Mike

Scharf, in the past 2 years (especially the last 6 months) than can probably not be repaid in the

next three decades. All have provided tremendous input not only on this current research, but on

future work as well in both science and in life. All of which, is greatly appreciated and I

sincerely hope that their input will not end upon graduation.

Although I recently met Dr. Roberto Pereira, his influence will last the rest of my life. His

drive for science and his skill at teaching is inspiring and his support has been nothing less than

remarkable. He would always find time to listen, critique, review, and analyze anything I

brought to him despite his own hectic schedule and workload.

My research would not have even begun without the tremendous work by the Urban

Entomology Laboratory Fly Group and the generosity of Dr. Phil Kaufman for allowing the fly

group to use his laboratory space. Ricky Vazquez orientated me and gave me the tools needed to

continue his successful fly program. Dr. Matt Aubuchon, Ryan Welch, Terry Krueger, and Mark









Mitola made the fly program function every day over the last two years and without all of their

consistent work the colony would have surely crashed. Terry Krueger provided more help than

any other when it came to my research. He was always there to provide a helping hand and

would often do recordings when I was unavailable to do so. Without his help, this research

could not have been completed.

Two fraternal brothers helped in every way they could throughout my University of

Florida experience. Tiny Willis kept me completely stocked and ready to go at all times and was

always a pleasure to talk to about life, politics, and the good ol' days whenever I needed a break

from my research, writing, or class. HM1 (SCW) Joseph W. Diclaro II, is an old friend who

seems to follow me wherever I go and I appreciate that because it seems good things come

whenever he does. Hopefully his time at the University of Florida will be as memorable as mine.

So mote it be.

Many new friends have been made and their friendship is valued. To mention them all

would cumbersome. The entire urban entomology lab and Urban Entomological Society have all

been great to work with a truly extraordinary group of scientists. In addition, the whole

administrative staff in the Entomology Department has been outstanding. If it wasn't for them I

can say with complete confidence that my graduation, as well as many others, would not come to

pass.

Finally, my family has been the foundation for this entire chapter in our lives together; my

wife, Karina, the cornerstone. Never once did she question time allotted away from her or our

two children, Conrad and Kyra, for or was she ever not willing to edit manuscripts when

obviously the subject matter did not suit her interests. Conrad and Kyra deserve a very, very

special thank you for being such really great children always Daddy's little helpers.










TABLE OF CONTENTS

page

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

L IST O F TA B LE S ......... .... ........................................................................... 8

LIST O F FIG U RE S ................................................................. 9

ABSTRAC T ................................................. ............... 10

CHAPTER

1 STATEMENT OF PURPOSE ................................... 12

2 R EV IEW O F LITER A TU R E ............................................... ......................... ...............14

Classification, Origin, and Distribution ............................................. 14
Id e n tific a tio n ..................................................................................................................... 1 4
aEgga ......( gg .................... ....................................... 14
Larva (maggot) ..................................14
P u p a ................... ...................1...................5..........
A d u lt ................... ...................1...................5..........
Sex D differentiation ................................................................ 16
L ife C y c le ............... ... ... ............................................................................... ....... 1 6
Nutrition, Longevity, and Overwintering ........................................... .. ................17
Flight, M ovem ent, and Resting Behavior............................................... 19
P est Status and H health Im portance ................................................................................... 2 1
C o n tro l .............. ...... .............................................................2 3

3 INSECTICIDE-IMPREGNATED CORDS FOR HOUSE FLY CONTROL ........................29

In tro d u ctio n ............................ ................................................................................................. 2 9
M materials an d M eth o d s ...........................................................................................................3 0
In sects. ......................................................................... 30
Laboratory A renas. ................................................................. 31
Cord Attractiveness Bioassay ................ ........ ...... ........31
Impregnated-Cord Laboratory Bioassays .............. ................................32
Impregnated-Cord Field Cage Bioassay................................ ................. 33
Statistical A analysis. ................................................................34
R e su lts ................... ...................3...................5..........
D iscu ssio n ................... ...................3...................7..........

4 EVALUATION OF A NEW IMIDACLOPRID BAIT FOR HOUSE FLY CONTROL ......46

In tro d u c tio n .............. ............... ....................................................................................... 4 6
M materials and M methods ...................... ................................. 48


6









In sects .............................................................................................................................4 8
L laboratory A rena D design. ...................................................................... ...................48
Field C age D design. ................................................................49
Fly Bait Com prisons. ...................... ............................................. .....49
B ait-Treated C words. ....................... ...................... .. .. ........... .... .. .....51
D ata A n a ly sis ...................................................................................................................5 2
R e su lts ................... ...................5...................2..........
Fly B ait Com prisons. .................................................. .... ...... .... ....... 52
B ait-Treated C words. ....................... ...................... .. .. ........... .... ....... 53
D iscu ssio n ................... ...................5...................4..........

5 SUM M ARY AND CON CLU SION S....................................................................... ... ..... 64

APPENDIX

1 REVIEW OF INSECTICIDE-IMPREGNATED CORDS.................................... ............... 67

2 R E V IE W O F F L Y B A IT S ............ ............................... ...................................................70

3 REVIEW OF INSECTICIDES EVALUATED....................... ....... ...............72

F ip ro n il ................................................................................................................................... 7 2
In d o x acarb ..........................................................................7 3
Im id a c lo p rid ....................................................................................................7 4
M ethom yl ..................................................................................75

LIST OF REFERENCES .......... .... ................................ ............... 77

B IO G R A PH IC A L SK E T C H ................................................................................................... 85









LIST OF TABLES


Table page

3-1. Efficacy of various cords impregnated with 0.1% fipronil or 0.6% indoxacarb on
female house flies. .................................... .. ... ... .. ................. 40

3-2. Cumulative number of dead flies and percent fly count reduction in relation to
control fly counts of house flies exposed to 0.1% fipronil- and 1.2% indoxacarb-
im pregnated cords in field cages............................................... ............................. 41

4-1. Number of dead and percent fly count reduction in relation to control fly counts of
house flies exposed to imidacloprid bait-treated lattice squares in field cages. ................59

4-2. Number of dead and percent fly count reduction in relation to control fly counts of
house flies exposed to imidacloprid bait-treated cords in field cages. ...........................60









LIST OF FIGURES


Figure page

3-1. Laboratory and field experimental design elements.............. ............ ....................... 42

3-2. Attraction of female house flies to various natural and synthetic cords.........................43

3-3. Female house fly mortality exposed to various natural and synthetic cords treated
with 0.1% fipronil for 24 h (A) and 0.6% indoxacarb for 48 h (B). ...............................44

3-4. Comparison of the most attractive cord (manila) and the least attractive cord (nylon
parachute) in the cord attractiveness experiments. ................................. .................45

4-1. Mortality of female house flies exposed to imidacloprid and methomyl granular
scatter baits and a sprayable imidacloprid bait. ...................................... ............... 61

4-2. Morbidity (knockdown) of female house flies exposed to natural and synthetic cords
dipped in a 2.5% solution of imidacloprid sprayable bait. ..............................................62

4-3. Mortality of female house flies exposed to natural and synthetic cords dipped in a
2.5% solution of im idacloprid sprayable bait. ........................................ .....................63









Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science

POTENTIAL OF INSECTICIDE-TREATED CORDS AND SPRAYABLE BAITS FOR
CONTROL OF HOUSE FLIES (DIPTERA: MUSCIDAE)

By

Jeffrey Conrad Hertz

August 2007

Chair: P.G. Koehler
Major: Entomology and Nematology

House flies are often controlled using insecticides when the source of infestation can not be

located or remedied by non-chemical methods. Historically, house flies have shown a

tremendous potential to develop insecticide resistance and with few classes of insecticides

currently registered for house fly control, new products and methods need to be evaluated to

prevent future control failures. This research evaluated the potential use of two innovative

methods to control house flies: fipronil- and indoxacarb-impregnated cords and a sprayable

imidacloprid fly bait.

For the insecticide-impregnated cord studies, eight various natural and synthetic cords

were evaluated to determine which cords were attractive to house flies. Natural cords were more

attractive than synthetic cords; the plant-based manila cord was most attractive and the nylon

parachute cord was least attractive. The most attractive cords (manila, cotton, wool, nylon, and

polypropylene) were treated with 0.1% fipronil or 0.6% indoxacarb and evaluated in the

laboratory to determine their effectiveness. All cords were more effective than the impregnated

cotton cord except the fipronil-impregnated nylon cord (LT90) and the indoxacarb-impregnated

polypropylene cord. The wool cord was the most effective, LT50 (Fipronil = 12.9 h; Indoxacarb

= 32.6 h) and LT90 (Fipronil = 22.4 h; Indoxacarb = 51.5 h). The wool cords were impregnated









with 0.1% fipronil and 1.2% indoxacarb and evaluated in a controlled field environment with

fresh cords and cords that were aged 4 wk. No significant differences were seen between fly

count reductions of either treatment. Both treatments reduced fly counts by >57% by 24 h and

>87% by 48 h with both fresh and aged cords. A reduction in efficacy was seen with aged cords.

The new imidacloprid sprayable fly bait formulation was compared against two commonly

used dry scatter baits in the laboratory and against a granular imidacloprid paint-on bait in a

controlled field setting. Additionally, the sprayable bait was evaluated for use in impregnated

cords. No differences were seen in mortality between the three scatter baits in the laboratory or

between the imidacloprid baits in the field cages. Both imidacloprid baits reduced fly counts by

>70% in the field within 24 h, but were not effective after treatments were aged for 2 wk. When

various cords were treated with 2.5% of the new bait, the wool cord had higher mortality (74%)

compared to the other natural and synthetic cords tested. Knockdown recovery was observed

with all bait-treated cords in the laboratory, but was not determined to occur in the field cages.

The bait-treated cords reduced fly counts by >82% with fresh cords and cords aged for 4 wk.

Impregnated cords and the new sprayable bait should prove to be valuable tools in

established fly management programs in urban, agriculture, and military settings. Fipronil and

indoxacarb are not currently registered for house flies, but both appear to be effective

insecticides for their control.









CHAPTER 1
STATEMENT OF PURPOSE

Throughout history flies have undoubtedly been a nuisance to both man and animal alike;

however, because of their propensity to frequent pathogen-rich filth they do pose a human health

risk. House flies have been shown to transmit numerous pathogens and their synanthropic

behavior may make house flies one of the most troublesome insect vectors (West 1951,

Greenberg 1973). This is especially true in areas affected by natural disasters or conflict. Often

times following these chaotic events, basic sanitation measures are out prioritized for casualty

recovery and, as a result, tremendous populations of house flies emerge.

Chemical insecticides are often used in these situations or any situation where rapid house

fly control is needed. Today there are more chemicals registered as insecticides than ever before,

but few of these insecticides are registered for house fly control. The insecticides that are

registered for house fly control only come from five chemical classes: organophosphates,

carbamates, pyrethrins/-oids, triazines, and neonicotinoids. The organophosphates and the

carbamate insecticides continually get further restrictions by the Environmental Protection

Agency (EPA) limiting their use and their future availability in the United States may be bleak.

In addition, house flies have consistently shown the ability to develop resistance to all chemicals

used to kill them (Liu and Yue 2000, Scott et al. 2000, Kaufman et al. 2001) and having only 3-5

chemical classes to rotate with may prove to be detrimental to a fly management program. There

is an immediate need for new insecticides registered and new techniques for house fly control to

prevent future control failures.

In 2004, the Department of Defense (DOD) established the Deployed War-Fighter

Protection (DWFP) program to develop and test management tools for pest and vector species,

including house flies, which transmit diseases to the deployed war-fighters. The Armed Forces









Pest Management Board (AFPMB) administers the DWFP program and specifically requested

research to improve or develop integrated filth fly control strategies and non-conventional

pesticide methodologies. Insecticide-impregnated cords and sprayable fly bait may be beneficial

tools that the deployed war fighter could use for fly management programs. The research

contained herein was designed to provide new information regarding these techniques to the

DOD.









CHAPTER 2
REVIEW OF LITERATURE

Classification, Origin, and Distribution

The house fly, Musca domestic, belongs in the class Hexapoda, order Diptera, suborder

Brachycera, infraorder Muscomorpha (Cyclorrhapha), and family Muscidae (Triplehorn and

Johnson 2005). It was first described by Linnaeus (1758).

It is believed the family Muscidae evolved sometime during the Permian period of the

Paleozoic era (Lambrecht 1980). The exact origin may never be known, but many speculate that

the house fly originated in the Middle East area of the Palearctic region (Skidmore 1985, Pont

1991) and was distributed through multiple introductions into the New World (Marquez and

Krafsur 2002). Today, house flies are one of the most commonly found synanthropic pests. It is

found in virtually every region of the globe that man or animal exist. The only exception is

areas, such as high altitudes and the arctics, which are prone to extreme cold temperatures (West

1951).

Identification

Egg

House fly eggs are white, bluntly rounded, banana-shaped eggs approximately 1 mm in

length, and often laid in clusters (Keiding 1976). The egg widens in size posteriorly to anteriorly

and the dorsal surface has two longitudinal, curved ridges that narrow just prior to reaching the

caudal end.

Larva (maggot)

House flies have three larval instars. Each instar has no eyes, legs, antennae, or

appendages and are commonly known as maggots (Moon 2002). The maggot has a rounded

posterior that tapers to a point towards its head. A pair of black spiracular plates is located









posteriorly, which progressively becomes more chitinized and "D-shaped" through molts. First

and second stage larvae have two spiracular openings (slits) used for gas exchange and a third

opening appears on the third instar larvae (Moon 2002). Prothoracic spiracles are fan-shaped

and appear after the first molt. A cephalopharyngeal skeleton, comprised mainly of sclerotized

"mouth" hooks, is located at the anterior end of the larvae.

Pupa

House fly pupae are approximately 6.3 mm in length (West 1951). At the beginning of

pupation, the puparium is white in color but eventually becomes reddish-brown. The puparium

is medially enlarged with bluntly rounded ends. Two pupal horns are located laterally just prior

to the posterior boundary of the first abdominal segment (Siriwattanarungsee et al. 2005).

Posterior spiracles are located on the posterior end and appear as two flat, circular prominences.

The anterior spiracle is situated on the puparium in the same location as in the third instar larvae

(Siriwattanarungsee et al. 2005).

Adult

The adult house fly is a medium-sized (6-9 mm) gray insect with large brown compound

eyes (Moon 2002). On the vertex, between the eyes, lies the ocellar triangle containing the three

simple eyes. The house fly's antennae are also located between the eyes, within the triangular

facial depression. The antenna is six segmented, but only appears to be four. The first three

segments, the scape, pedicel, and large first flagellar segment, give rise to the three-segmented

arista. Segment one and two of the arista is ambiguous; segment three is bristlelike. The

sponging proboscis of the house fly terminates to a heart-shaped sucker. The proboscis can be

greater than the length of the head when fully extended or obscure when fully retracted. Two

brownish-black maxillary palpi lie on the anterior margin of the proboscis.









The thorax of the house fly has four black longitudinal stripes that can be viewed dorsally.

Attached laterally to the mesothorax are two membranous wings. The wings, when extended,

are approximately twice the distance of the fly's length. At rest, the house fly pulls the wings

back incompletely over the abdomen forming an overall triangular appearance from above. The

fourth longitudinal wing vein sharply angles towards the wing apex. Situated below each wing is

a knob-shaped organ used for equilibrium called the haltere. The legs of the house fly attach

ventrally to each segment of the thorax and all legs have five-segmented tarsi. The first tarsal

segment is much longer than all other segments and the fifth segment bears two claws, a hair-like

empodium, and a sticky pad called a pulvillus.

The abdomen is gray, dorsally, and cream-colored ventrally. Five pairs of spiracles line

the ventral surface of the female; six pairs line the ventral surface of the male. The tip of the

abdomen ends in either the sclerotized genitalia of the male or the retracted ovipositor of the

female.

Sex Differentiation

Adult female house flies are almost always larger than adult males. Additionally, males

can be differentiated from adult females by locating the dark sclerotized genitalia plate located

on the distal aspect of the abdomen. The tiny mark made by the ovipositor tip of the female is

very distinctive compared to the male genitalia especially when females are gravid.

Furthermore, adult house flies can be separated by the gap distance that divides the compound

eyes. Females have a much wider space separating the eyes when compared to male

counterparts. No differentiation can be made in the immature stages.

Life Cycle

Male and female house flies can successfully mate 24 hours after emergence from the

pupae (Murvosh et al. 1964). Prior to copulation, a male will seize a resting female or strike a









flying female at which point they fall to a surface. If a copulating pair is disturbed while mating

they may attempt to fly a short distance to an alternate surface. Copulation can last for more

than 1 h, but sufficient sperm transfer can occur in less than 10 min (Murvosh et al. 1964). Once

successful copulation takes place, the female is fertilized for life. Batches of up to 150 eggs are

laid 4-8 days after copulation (West 1951). The female house fly carefully embeds her eggs into

practically any fermenting organic material. The eggs hatch within 24 hours, and 1st instar larvae

emerge and begin to feed (West 1951). The larvae undergo two molts within 3-5 days before

pupation (Hogsette 1995). Pupation begins when the 3rd instar larva stops feeding and constricts

within its own integument. This makes a white puparium which turns reddish-brown within 24

hours. After 3-5 days, the adult breaks through the anterior end of the puparium using a

temporary, inflated sac located on its head called the ptilinum. Once free from the puparium, the

newly emerged adult house fly hops around to let its wings extend and cuticle harden.

Nutrition, Longevity, and Overwintering

House flies larvae have been reared in the laboratory on practically every type of filth

imaginable. Today, they are most often reared in a medium containing animal feed and water

(Hogsette 1992). Fermenting odors attract gravid females to oviposit on breeding sites in the

field, but understanding precisely what nourishes a maggot within the medium is not fully

understood. All house fly maggots are saprophagous and feed on liquids or substrates that are

readily dissolved by droplet regurgitation (Nation 2002). It was originally thought that bacteria

were essential in the development of house fly maggots, however many have successfully reared

them in aseptic media (Brookes 1956, Monroe 1962). Despite this, bacteria still may have

provide nutritional value (e.g. vitamins) to maggots (Zurek et al. 2000).

Adult house flies are omnivorious and emerge with little stored energy and nutrients

(Moon 2002). They begin to feed within 2-24 hours after emergence (Keiding 1976). In order to









survive, they must find a sugar source, or other assimilable starch, and water (West 1951). In

addition, female house flies require a protein source for vitellogenesis.

When feeding, adult house flies are attracted first visually, then when they are within a

detectable range, by smell using their antennae (Keiding 1976). Flies locate the source of the

aroma by smelling the substrate with chemoreceptors located on the lateral aspect of their 2-5

tarsi. Once their tarsi are in contact with a suitable substance, the fly extends its proboscis and

begins to feed. Liquid substances can be readily imbibed, but solids are ground down using the

prestomal teeth on the proboscis and emulsified using a vomit drop originating in the crop and

salivary glands. The largest particle a house fly can ingest is 40 [t (Greenberg 1973). Ingested

liquid and emulsified particles enter the pseudotracheae and then pharynx. Once past the

pharynx, liquids pass into the crop and emulsified food particles enter the proventriculus then the

ventriculus. The crop is connected to the pharynx by a long slender tube lined with numerous

sphincters that controls the flow of liquid back to the abdomen where the bilobed crop is housed.

The heamolymph osmotic level dictates the rate the crop empties into the ventriculus. The more

concentrated the sugar meal, the slower the crop empties (Greenberg 1973). The ventriculus

empties into the longest part of the alimentary track, the proximal intestine. The proximal

intestine is divided from the distal intestine by excretory organs called the Malpighian tubules.

The distal intestine terminates at the anus.

Longevity of any organism can be highly variable. Food availability, environmental

conditions, and activity of an individual fly greatly influences how long it will live. Of the three

survival-mandated nutrients, sugar is the most critical for survival. Lysyk found that the

availability of sucrose was the most important factor promoting longevity, followed by other

food sources (manure, milk) and temperature (1991). Flies will live 50% longer on sucrose









alone, than they do on water alone (Greenberg 1960). However, flies without water generally die

within 48 hours (West 1951).

Temperature is inversely proportional to the life span of the adult house fly; higher

temperatures reduce the life expectancy, while lower temperatures increase it (West 1951). In

laboratory conditions where adequate food is provided ad libitum and environmental conditions

are controlled, male house flies can live up to 40 days and female house flies can live up to 60

days (Rockstein 1957). In the field, house flies are estimated to only live about 10 days

(Hogsette 1995). Bucan and Sohal (1981) found that adult males and females isolated from the

opposite sex live longer than when they are housed together.

To increase their survivability when temperatures drop below optimum levels, house flies

survive by overwintering in buildings and animal confinements. All life stages are susceptible to

subzero temperatures, so microclimates must exist that allow flies to propagate. Rosales et al.

(1994) concluded that house flies require habitats that are above -5C, and must stay above 10C

long enough for the house fly to complete its life cycle.

Flight, Movement, and Resting Behavior

Flies have two wings located on the lateral aspect of the body on the pteropleura. Directly

above the metathoracic coxae are the vestigial wings, or halteres, which are used as gyroscopes

for equilibrium. Like other flying insects, a house fly achieves flight by creating wing

movement through indirect thorax compression and decompression caused by the flight muscles.

These flight muscles comprise approximately 11% of the total body weight in the genus Musca

(Greenberg 1973). This musculature makes house flies extremely strong fliers and very capable

of flying upwind in mild and moderate winds.

Fly movement can be classified as dispersal, dispersion, or migration (Greenberg 1973).

Dispersal is any active movement within a relatively small defined area. House fly problems are









often localized near a source of infestation (Howard 2001, Nazni et al. 2005). Often times,

dispersal will be dependant on the sun light. Flies tend to follow the sun and more will be

located where the sun is shining (Anderson 1964). This is especially true in cooler temperatures;

in hot temperatures, flies may avoid the sun and search for cooler locations. Dispersion is the

movement of flies between adjacent areas and often involves the movement assisted by passive

transport. Passive transport can occur on garbage trucks, tractors, automobiles, or any other

vehicle including strong winds. This is often seen when breeding sources are sporadic or when

no breeding sources are near and flies move into the area in search of new oviposition sites. This

movement is why flies are found in areas where no apparent fly breeding material is present.

Migration is any directed and sustained flight that often occurs seasonally. This type of

movement is not normally associated with house flies, however, many have been trapped in areas

that would suggest that migration was the only possible explanation (West 1951, Jones et al.

1999). Passive transport may also play a large role in these situations.

Fly movement can be influenced by many factors such as odors, wind, weather, time of

day, and population structure. Food and oviposition sites are probably the most critical factors

(Bishopp and Laake 1921). However, many questions still need to be answered on what is

attractive or needed by house flies since flies will often leave one oviposition site in search for an

alternative site although sources are immediately available and sufficient for survival. When

seeking these new food sources or oviposition sites, flies can fly upwind in mild to moderate

wind speeds, but strong winds, or even shifting winds, can disperse house flies to areas where

survival may not be suitable. Taylor (1974) found that house flies are day fliers and their flight

activity increases with high temperatures (25-30C) and low humidity (50-65%). Flies always









seek temperatures above 15.5C, but are capable of flight at temperatures below 55C (Greenberg

1973).

House flies have distinct resting behavior. In warmer climates, flies prefer to rest at night

outdoors on low hanging twigs of trees and bushes, but may still be seen resting indoors on wires

or cords close to the ceiling (Scudder 1949, Keiding 1965). In colder climates, house flies will

rest exclusively indoors. In all cases, flies tend to rest on objects with distinct edges less than 4.5

m from the ground, shielded from direct wind (Scudder 1949). Keiding and Hannine (1964)

found a distinct preference for house flies to rest on objects suspended vertically from ceilings,

however, Fay and Lindquist (1954) found no differences in orientation of suspended cords.

The visual orientation of house flies to objects has been widely disputed. Objects that are

light in color, smooth, or metallic are highly avoided by flies; whereas, objects that are dark in

color and rough are generally more frequently rested upon by flies (Arevad 1964). Hecht et al.

(1968) performed a number of indoor and outdoor experiments to determine the attraction of

house flies to different colored cardboards. They found that black was most preferred indoor and

white was the most preferred outdoor. When combining the indoor and outdoor results, the red

colored surfaces were most preferred. The least preferred colors were blue (indoor) and brown

(outdoor). The attraction to the white surface outdoors was attributed to a fly's attraction to

ultraviolet light because of the reflective qualities of the white cardboard. Flies see wavelengths

between 350-480 mut (McCann and Arnett 1972). Contrasts of colors (dark on light/light on

dark) may be very important to the attraction of house flies. Howard and Wall (1998) counted

more flies on white surfaces with black backgrounds than on any other black/white combination.

Pest Status and Health Importance

House flies are renowned for their ability to annoy anyone and anything they are near. It

only takes one house fly to turn a customer away from a restaurant and only a few to disrupt









production and morale at a work site. Most see house flies as a sign of unhygienic conditions

and attempt to avoid them at all costs. Flies leave fecal and vomit spots on work equipment,

consumables, and personal items most of which are not be generally quantified in economic

losses but their impact can easily be seen when comparing two similar establishments one with a

fly problem and the other lacking a problem. Litigation cases due to house flies have increased

in the United States recently due to the migration of urban dwellers deeper into rural settings

where livestock and poultry farms have a relatively large abundance of flies a situation many

urban dwellers may not be familiar with.

Although primarily nuisance pests, house flies do pose a risk to the health and well-being

of man and livestock. Because of a house fly's behavior and survival needs, it frequently comes

in contact with pathogenic organisms. House flies are extremely capable of transmitting

pathogens mechanically (West 1951). West (1951), and most recently Greenberg (1973), have

compiled extensive lists of the pathogens (bacteria, viruses, fungi, protozoa, and nematodes) the

house fly is capable of transmitting. The transmission of Campylobacter spp., E. coli 0157:H7,

H. pylori, C. parvum, and G. lamblia are probably the most significant pathogens capable of

being transmitted by the house fly recently reported (Shane et al. 1985, Grubel et al. 1997,

Kobayashi et al. 1999, Graczyk et al. 2003).

Of particular importance is the exponential proliferation of house flies following situations

arising from natural disasters or conflict. Following the tsunami that devastated parts of

Indonesia in 2004, sewage and drainage systems were destroyed leaving sewage pools that bred

numerous species of filth flies (Burrus 2005). This is often what occurs following any of these

chaotic events; communities are left in shambles and multiple oviposition sites develop due

infrastructural collapse or basic municipal sanitation being out prioritized. Large fly populations









then develop and epidemic levels of diarrheal cases normally follow (Thornton et al. 2005,

Watson et al. 2007). This reduces military readiness and stresses health care systems (Putnam et

al. 2006). The direct impact of house flies on disease transmission in these situations is either

not measured or often not measurable because the same pathogens transferred by house flies can

be just as easily transferred by man or other organisms.

Control

The house fly is best controlled through a fly management program based on the sound

principles of Integrated Pest Management (IPM), including a combination of monitoring,

cultural, biological, and chemical control measures (West 1951, Keiding 1976). Monitoring is

any technique employed to determine presence/absence and peak/trough flows of house fly

populations. Cultural controls are any measures that deliberately alter the life cycle of the house

fly without the use of chemical or biological agents. Biological controls are agents or organisms

that alter the life cycle of the house fly and chemical controls are naturally- or synthetically-

derived chemicals that can alter the life cycle of the house fly.

Every aspect of the fly management program should have some form of fly monitoring to

determine when and what type of approach should be employed (pre-treatment survey) and to

see the effectiveness of the treatment (post-treatment survey) (Keiding 1976). In the pre-

treatment survey, house fly density, distribution, and behavior should be noted to help determine

which treatment option to use. Post-treatment surveys normally only need to monitor fly

densities unless failure occurred. In this case, a complete reassessment should be done to

include some form of monitoring for insecticide resistance.

There are four basic methods to obtain a fly population index: counting flies, counting fly

specks, netting flies, or trapping flies (Keiding 1976). If counting adult flies, several methods

can be used. The classic technique is the use of a Scudder grill a simple grid constructed of









wood that can be placed over an infestation source and then all flies landing on the grid are

counted (1947). A similar technique that can be used in practically any situation is to simply

mark an area (preferably near infestation or resting areas) and count flies landing on it. Counting

fly specks left on index cards may be the method of choice today for indoor sampling because of

its simplicity. Spot cards can be positioned in standard locations throughout the infested area

and will give a good representation of the fly populations over time. Simply hang them and

check back on them after a designated time period. Over time it will show peak and troughs of

fly populations; in addition, the spot cards can be archived and marked with any insecticide

treatment used to provide additional documentation for resistance monitoring. Some users,

however, prefer to use destructive sampling. In these cases, baited traps, sticky ribbons, or even

netting flies can work well.

All of the above methods work and can give consistent numbers indoors as long as the

same sampling method is used for all counts. However, infestations that occur outdoors are not

as easily monitored. With outdoor sampling, spot cards are an unrealistic method because

placement would be difficult and precipitation would destroy them. Scudder grills and other fly

count techniques are subject to wide variation depending on positional effects, time of day, and

weather conditions (Geden 2005). Baited traps and net sweeping may be the best techniques to

use outdoors for surveillance work, but these methods are destructive and not suitable in every

situation. Beck and Turner (1985) found that using a simple visual index correlated better with

absolute fly densities than spot cards, sticky ribbons, scudder grill and fly counts, but these

indexes are very subjective and will vary between persons making the counts. Perhaps, the best

way to monitor house flies outdoors is to use a combination of the methods described.









By far the most important aspect of a fly management program is the use of cultural

controls. Cultural controls target the breeding and feeding sites of adult and larval flies.

Additionally, they are used to prevent adult flies from contacting food, pathogens, and man

(Keiding 1976). Garbage is the main source of infestation in an urban environment and manure

is the main source in an agriculture environment, however, both sources can be found in any

environment. In developed urban communities, these breeding sites are normally controlled by

very established municipal sanitation measures (e.g., closed sewers, garbage removal, etc.)

(Hogsette 1995). Agriculture facilities have to physically remove manure or bake it by covering

dung heaps with plastic sheeting. Garbage should be removed from the area at least twice

weekly or burned (Keiding 1976). Windows, screens, and doors should all be in good repair and

kept closed to prevent flies from entering establishments. The installation of air curtains on

doors and windows that are frequently opened and closed is an energy efficient option that can

reduce the number of flies that enter.

Traditionally baited traps, light traps, electrocuting light traps, and sticky ribbons have

been used as cultural control measures, but their effectiveness at reducing house fly populations

are limited and their use should primarily be considered as a monitoring technique. However,

these methods do trap and kill flies so using them should never be automatically ruled out. In

fact, if the likelihood of large infestations does not exist, then traps are a good method for killing

flies (such as in grocery stores and restaurants) but some considerations should be made prior to

their use. Baited traps generally can not be used indoors or near residences because the odor

associated with this method is repulsive to humans (Pickens et al. 1973). Electrocuting light

traps release fly parts, bacteria, and viruses and may be just as unhygienic as the fly itself and

should not be used in areas were conditions need to remain relatively aseptic (Urban and Broce









2000). Lighted traps need bulb replacement approximately every six months, and sticky ribbons

need replaced frequently due to dust and fly cadaver build-up. Flies that do make their way into

an establishment can be physically removed by numerous devices, but the most common

physical cultural control method is the good 'ol fashioned flyswatter. A novelty gadget used for

killing insects, including flies, has recently sparked numerous videos on the World Wide Web.

The device is a combination electrocuting trap and fly swatter. This device may be interesting

and fun, but the same risk is associated with it as the regular electrocuting light traps.

The second most important principle of a good fly management program is the use of

biological controls. Every stage of a fly's life cycle is vulnerable to attack by some form of

biological control (West 1951). The eggs are often predated upon by mites, earwigs, ants, and

some beetles. Larvae are also attacked by mites, earwigs, and beetles, as well as some birds,

wasps, and other Dipteran larvae. Pupae are often parasitized by small wasps and some beetles,

others can be eaten by birds and large beetles. Many adult house flies meet their demise thanks

to predatory insects mantidss, flies, dragonflies, wasps, ants) and arachnids. Many other adults

are eaten by reptiles, amphibians, small mammals, and birds. House flies are also prone to

infections by bacteria and fungi. Fortunately, all of these biological controls are already

abundant in a fly's natural environment and the goal of a fly management program should be to

maintain or supplement these existing populations (Geden 1995). Parasitic wasps can now be

purchased commercially and their successful use is variable (Axtel 1999).

The use of chemical insecticides is the third component of a good fly management

program. Chemical insecticides provide quick results (i.e. dead flies) and satisfactory control in

one or two days, but their use should be limited to reduce the likelihood of resistance evolution

and maintenance of non-target organisms. Chemical insecticides can be applied several different









ways: residual surface treatments, larviciding, space sprays (including aerial spraying), and baits.

The most common method for fly control is the use of space sprays and dry insecticidal baits.

Residual surface treatments can be applied to any surface in any location the label allows

but they are most effective when applied directly to fly resting areas (Keiding 1976). Neglecting

to monitor and treat areas where flies are primarily resting will result in excess insecticide usage

and population reductions of non-target organisms. Several insecticides are available for

residual treatments; most are organophosphate based. All are generally good for long term

control, but there excessive use may increase selection for insecticide resistance. One technique

for residual insecticide application is the use of insecticide-impregnated cords which target the

distinctive behavior of flies to rest on objects with edges (Scudder 1949, Keiding 1965). This

method is thought to be less likely to select for resistance because the treated area is small and

treatments can be readily removed or replaced with additional cords treated with insecticides

from different chemical classes (Appendix A).

Larviciding with insecticides sounds great in theory because larvae are relatively non-

mobile compared to the adult flies, which have the ability to fly to different areas if one is found

unsuitable; however, larviciding is really not a practical method for extended control. Larvicides

have to be applied frequently because they are applied to areas such as garbage and manure -

both of which constantly accumulate. Larvicides kill non-target organisms coming in to feed on

the house fly immature stages and would reduce those populations over time. In addition, if the

same class of insecticide is used for larviciding that is used for adult control resistance selection

would be rapid. Larvicides are effective if sites to be treated are expected to exist for a short

period of time; in these instances several organophosphate insecticides and insect growth

regulators can be used.









Space sprays are primarily pyrethrin- or pyrethroid-based insecticides, but some may also

be organophosphate-based, that are sprayed into the air in a fly infested space or over a fly

infested area. These types of insecticides target the fly nervous system and cause rapid

knockdown of contacted flies (Yu 2007). Space sprays are more effective when an abundance of

flies are concentrated in one area; this occurs mainly in the evening indoors and in the morning

outdoors (Keiding 1976). They have little to no residual and have to be reapplied frequently

(daily) in areas of large infestations. Another type of space treatment is made through the use of

insecticide vaporizers. The only vaporizer currently available is formulated with the

organophosphate dichlorvos, but its use is becoming more restricted and its future longevity may

be short lived.

Insecticide baits are easy-to-use insecticides that have added attractants into the

formulation matrix to draw flies into the treated area to contact the insecticide either by ingestion

or contact. A basic bait matrix is a simple solution of sugar, water, and an insecticide. Complex

bait matrices contain multiple sugars, pheromones (Z-9-tricosene), and other substances found

attractive to house flies. The most widely used fly baits available are formulated in dry granules

as scatter baits containing carbamates and neonicotinoids (Appendix B), however, other bait

products are available and frequently used. Like space sprays and larvicides, baits have to be

frequently applied because environmental conditions degrade them or they become covered by

manure or garbage. Also, when baits are used in areas with large fly populations, the flies will

consume the bait rapidly leaving little bait behind for the immature stages that will eventually

emerge.









CHAPTER 3
INSECTICIDE-IMPREGNATED CORDS FOR HOUSE FLY CONTROL

Introduction

The house fly, Musca domestic L., is widely considered the most common nuisance pest.

Their nuisance pest status can quickly change to a public health risk if fly populations occur near

inhabited areas where pathogen-rich oviposition sites are found. Areas stressed due to natural

disasters, humanitarian crises, or combat are often plagued by large fly populations (Rosales and

Prendergast 2000, Burrus 2005, Thornton et al. 2005). The most effective way to control house

flies and reduce the risk of disease transmission is by eliminating their pathogen-rich oviposition

sites. Oviposition site removal may be impractical, especially in areas affected by natural

disasters and combat, where the oviposition sites are too numerous or difficult to reach.

The best control method to use when sanitation fails or when fly populations need to be

rapidly controlled are chemical insecticides. Chemical insecticides provide rapid kill of house

flies and markedly reduced fly densities can be achieved in as little as 1-2 days. Baits and space

sprays are the primary chemical insecticide methods used for house fly control today, but both

methods provide little or no residual control, and resistance to their active ingredients is well

documented in house flies (Georghiou and Lagunes-Tejeda 1991, Liu and Yue 2000, Scott et al.

2000). In addition, the number of registered insecticides available for house fly control in the

United States continues to decrease (Kaufman et al. 2001). New insecticides and application

methods are clearly needed to avoid future insecticide resistance problems.

Insecticide-impregnated cords have been used with great success to control flies and are

considered less likely to select for resistance than traditional residual sprays (Keiding 1976).

Their first use was in 1947 (Baker et al.) and by the mid-1950's, insecticide-impregnated cords

were commercially available and widely used (Fehn 1958, Smith 1958). The commercially









available cords contained 13.79% parathion and 3.54% diazinon (Smith 1958). Cords

impregnated with high concentrations (up to 25% active ingredient) of other organophosphate

and organochlorine insecticides were also widely used with great success (Kilpatrick and Schoof

1959, Keiding 1976, Rabari and Patel 1976). These products are no longer used today due to the

popularity of insecticidal baits and space sprays and because the Environmental Protection

Agency, acting under federal legislation, eliminated the use of their active ingredients.

The objective of this study was to investigate if cords impregnated with newer insecticides

would be an effective tool for house fly control. Specifically, the objectives were to: 1)

determine the attractiveness of various natural and synthetic cords to house flies, 2) determine

the effectiveness of fipronil and indoxacarb on the most attractive cord materials, and 3) evaluate

the effectiveness of the best cord/treatment combination in a simulated field environment.

Materials and Methods

Insects. The Horse Teaching Unit (HTU) strain of house flies, M. domestic L., reared at

the University of Florida in Gainesville was used for all experiments. Larvae were reared on a

diet medium, modified from Hogsette (1992), containing 3 liters wheat bran, 15 ml methyl

paraben, 1.5 liters water, and approximately 200 g (250 ml) dairy calf feed (Calf Manna

pellets, Manna Pro Corp., St. Louis, MO). All developmental stages were held at 26 + 1IC and

55% RH with a 12:12 (L:D) photoperiod. Adult flies emerged within screened rearing cages and

were provided granulated sugar, powdered milk, and water ad libitum.

For all assays, adult house flies (3-5 d old) were aspirated from the screened rearing cages

using a handheld vacuum with a modified crevice tool attachment. Flies used for the laboratory

assays were placed into a 50C environment for 5 min to subdue activity. Flies were then placed

on a chilled aluminum tray, sexed, and counted. House flies used for field cage assays were not









anesthetized, but were aspirated from the screened rearing cages and released directly into field

cages.

Laboratory Arenas. Arenas (31 x 25 x 21 cm) were constructed using PVC pipe (1.27

cm [0.5 in]) (Figure 3-1A). Rubber bands were used to establish individual treatment positions;

four treatment positions were used in the cord attractiveness bioassay and five positions were

used in the impregnated cord bioassay. All cords were attached to the treatment positions

vertically using paper clips and were uniformly distributed along the length of the arena. The

cord attractiveness bioassay held two randomly assigned cords at each treatment position and the

impregnated cord bioassay held only one cord at each treatment position according to a 5 x 5

Latin square configuration. Arenas were enclosed with a transparent plastic bag (3716 cm2 [24 x

24 in], 1 mil poly, Uline, Waukegan, IL).

Cord Attractiveness Bioassay. Eight cords were evaluated: nylon (Braided, Multi-

Purpose Braid 75 lb. load limit, Wellington Cordage LLC, Madison, GA), polypropylene

(Braided, Multi-Purpose Rope 56 lb. load limit, Wellington Cordage LLC, Madison, GA),

cotton (Braided, Multi-Purpose Sash Cord 28 lb. load limit, Wellington Cordage LLC,

Madison, GA), cotton wick (Sterilized roll, #200209, Richmond Dental Company, Charlotte,

NC), manila (Twisted, Natural Rope 108 lb. load limit, Wellington Cordage LLC, Madison,

GA), wool (Twisted, Natural Cord, Wooded Hamlet Designs, Greencastle, PA), leather (Tan

laces, #6192, Rothco, Ronkonkoma, NY), and parachute cord (550 test, white, purchased locally

from M & C Army Surplus Store, Gainesville, FL).

Fifty female flies were released into each arenas and 10% sugar water was provided ad

libitum. Number of flies resting on cords was counted every 10 min for 2 hr. Arenas were

lightly shaken between each count to displace flies from their resting positions. Four replications









were performed in the laboratory (28 1C) under continuous light on separate days using

different flies.

Impregnated-Cord Laboratory Bioassays. Cotton, manila, wool, polypropylene, and

nylon cords were selected from the cord attractiveness experiments to be evaluated in the

impregnated-cord experiments. Each impregnated-cord experiment consisted of 6 arenas. Five

arenas were organized into a 5 x 5 Latin square design, blocking for treatment position, and a

sixth arena was used as a control. The control arena had no treated cords and all cords within it

maintained the same cord positions throughout all experiments (left to right: position 1 = cotton;

2 = wool; 3 = manila; 4 = polypropylene; 5 = nylon).

Separate experiments were done to evaluate cords (15.24 cm length [6 in], 0.6 cm [0.25]

diam) impregnated with a 0.1% fipronil or a 0.6% indoxacarb solution. The 0.1% fipronil

solution was prepared by combining 2.7 ml of the formulated insecticide (Termidor SC, 9.1%

a.i., BASF, Research Triangle Park, NC) with 250 ml of tap water. The 0.6% indoxacarb

solution was prepared by combining 5 g of formulated insecticide (DPX MP062, 30WG,

DuPont, Wilmington, DE) with 250 ml of tap water. Cords were impregnated by dipping for -2

sec in the insecticide solution and were then allowed to dry in a fume hood.

Groups of 50 female flies were placed within each arena and provided a 10% sugar water

solution ad libitum. Mortality counts were recorded until at least 80% mortality was observed.

Due to the differences in the mode of action of the insecticides, mortality for flies exposed to

fipronil-impregnated cords was defined as the inability to remain standing; flies exposed to

indoxacarb-impregnated cords were considered dead if they were unresponsive to touch. Each

experiment was run in the laboratory (28 1C) under continuous light and replicated twice.









Impregnated-Cord Field Cage Bioassay. Cages (1.8 x 3.7 x 1.8 m) were constructed

from PVC pipe (2.54 cm [1 in] diam) and enclosed with mesh screening (Outdoor Cage,

#1412A, 18 x 14 mesh, Bioquip, Rancho Dominguez, CA). Black plastic sheeting (6 mil) was

used to line the floor. A sampling stage, constructed of two vertical cinder blocks and an

inverted storage bin (Palletote #1721, 37 liter, Rubbermaid, Winchester, VA), was placed in the

center of the cage (Figure 3-1B). On top of the sampling stage there were two 994-ml (1 qt)

chick waterers, one filled with 10% sugar water and the other with tap water, and a 60-ml plastic

cup filled with 8 g of previously used larval house fly medium. The chick waterers provided

enough sustenance for the duration of the test and the plastic cup was used as an attractant. The

plastic cup was covered with a paper towel and sealed with a rubber band to prevent flies from

ovipositing on the medium.

Treatments consisted of two long (0.9 m) and eight short (0.6 m) lengths of 0.1% fipronil-

and 1.2% indoxacarb-impregnated wool cords. The 0.1% fipronil solution was prepared by

combining 7.7 ml of the formulated insecticide (Termidor SC, 9.1% a.i., BASF, Research

Triangle Park, NC) with 700 ml of tap water. The 1.2% indoxacarb solution was prepared by

combining 28 g of formulated insecticide (DPX MP062, 30WG, DuPont, Wilmington, DE) with

700 ml of tap water. Each cord was treated in the same manner as the laboratory experiments,

except the cords were dipped and soaked for 1 min prior to drying.

Depending on fly availability, 27.5 35 ml (9.8 + 1.8 flies/ml) of flies was released into

each cage. After a 1-h acclimation period, pre-treatment fly counts were taken. Before fly

counts were taken, the operator walked three laps around the interior of the cage to disturb flies

from their resting positions and to recover any dead flies from the cage floor. Four consecutive

fly counts were then taken from the outside of the cage 1 min after exiting. All flies that landed









on the sampling stage, chick waterers, and plastic cup attractant were counted. Treatments were

then hung vertically from the mesh ceiling using paper clips in specific locations (Figure 3-1C)

and post-treatment fly counts were taken at 24 and 48 h using the same method described above.

After the initial 48 h evaluation, treatments were aged in the elements for four weeks, at which

point residual effectiveness was re-evaluated as described above. Three replicates were

performed at each treatment age (0 and 4 wk).

Statistical Analysis.

All statistical analyses were performed using JMP IN (SAS Institute 2005), except probit

analysis estimates were performed using SAS (SAS Institute 2001). For the cord attractiveness

experiments, the mean number of flies/cord was analyzed using a one-way analysis of variance

and contrasts were performed between natural and synthetic cords and the animal- and plant-

based cords. For the laboratory insecticide-impregnated cord laboratory experiments, mortality

data were corrected using Abbott's formula (1925) and arcsine square root-transformed. A two-

way analysis of variance was performed on the 24-h fipronil data and the 48-h indoxacarb data to

determine if treatment position had an effect on mortality. LT50 values were estimated by probit-

analysis regression (Finney 1971). Potency ratios, using the cotton cord as the standard, were

performed using the method described in Robertson and Preisler (1991). Slopes, LT50 values,

and potency ratios were considered significantly different if the 95% confidence intervals did not

overlap. For the field cage experiments, percent fly count reductions were calculated from the

control fly counts. Fly count reductions and mortality data (number of dead flies recovered from

cage floor) were then analyzed for each treatment age (0 and 4 wk). All means were separated

using the Student's T or Student-Newman-Keuls test (a = 0.05).









Results

In the laboratory studies, all flies fully recovered from chilling after approximately 45 min

at which point the flies were dispersed throughout the entire arena. Flies were more attracted to

the manila cord, which had significantly more flies resting on it than any other cord (Figure 3-2).

No significant differences were seen between the other natural cords or between the synthetic

cords; however, all synthetic cords had significantly less flies resting on them than the natural

cords (F: 112.69, df = 368, P = <0.001) and the plant-based cords were more attractive than the

animal-based cords (F: 11.64, df = 368, P = <0.001). The least attractive cord was the nylon

parachute cord.

The laboratory design had no position or interaction effects for either fipronil (F: 1.05; df=

4; P = 0.3982, F: 0.8347; df = 20; P = 0.6583) or indoxacarb (F: 0.71; df = 4; P = 0.5906, F:

0.32; df = 20; P = 0.9955) in the insecticide-impregnated cord experiments. At the 24 h

(fipronil) and 48 h (indoxacarb) recordings, all impregnated-cords had significantly higher

mortality than the controls (Figure 3-3).

House flies suffered significantly higher mortality when exposed to the fipronil-

impregnated wool cord than any other fipronil-impregnated cord at 24 h (93%). The other

fipronil-impregnated natural cords had percent mortalities below 15%, with manila causing only

5% mortality at 24 h. No significant differences in mortality were seen between the fipronil-

impregnated nylon and polypropylene cords or the fipronil-impregnated cotton and manila cords

at 24 h.

The indoxacarb-impregnated wool cord caused significantly higher mortality (85%) than

any of the other cords except for the cotton cord at the 48 h recording. No significant differences

in mortality were seen between the synthetic indoxacarb-impregnated cords or between the

cotton and manila indoxacarb-impregnated cords. Significant differences in mortality were seen

35









between the wool and manila indoxacarb-impregnated cords. The indoxacarb-impregnated

nylon cord caused the lowest mortality at 48 h (47%).

Fipronil- and indoxacarb-impregnated cords efficacy results can be viewed in Table 3-1.

In general, the fipronil impregnated cords had lower LT50 and LT90 values than the indoxacarb-

impregnated cords. Among the fipronil-impregnated cords, the wool cord had the lowest LT50

and LT90 values and the impregnated cotton cord had the highest LT50 and LT90 values. The LT50

values for the synthetic cords were relatively low compared to the other cords, but the LT90

values were no different from the cotton cord. The manila cord LT50 value was the second

highest, but had the second lowest LT90 value behind the wool cord; it is important to note that it

also had the highest slope compared to the other cords. All cords were more effective than the

cotton cord except for the nylon cord's LT90 value.

All indoxacarb-impregnated cords had LT50 values >32 h and LT90 values >51 h. The

indoxacarb-impregnated wool cord had lower LT50 and LT90 values than all other indoxacarb-

impregnated cords except for the manila cord, which showed no significant differences in LT50

values. The indoxacarb-impregnated polypropylene and cotton cords each had LT50 values of 52

h and LT90 values >100 h, which were the highest values for the experiments. No differences in

LT values were observed between the manila and nylon cords. All cords were more effective

than the cotton cord except for the polypropylene cord's LT50 and LT90 values.

In the field cage experiments, no significant differences in fly count reductions occurred

between the treatments (Table 3-2). Both treatments had >57% fly count reductions by 24 h and

>87% by 48 h, independent of the treatment age. Dead flies were collected from all cages at

every recording; significantly more were collected from the treatment cages than the controls.

Fipronil treatments had significantly more dead flies than the indoxacarb treatments with fresh









cords at 24 and 48 h and with aged cords at 24 h. No significant differences were seen between

the number of dead flies collected from the fipronil treatments and indoxacarb treatments at the 4

wk, 48-h recording.

Discussion

Insecticide-impregnated fly cords are based on a fundamental component in a fly's

behavior flies prefer to rest on objects with distinct edges, such as twigs, wires, cord, and line

(Scudder 1949). Since insecticide-impregnated cords only represent a small proportion of

available resting surfaces available to flies, it is assumed that factors which enhance a fly's

attraction to the cords would be beneficial to the effectiveness of the treatment. Surfaces which

are more attractive to flies would be expected to cause quicker mortality because of increased

exposure to the insecticide. Arevad (1965) found flies to favor dark, rough surfaces over light,

smooth surfaces. Specific factors influencing a fly's attraction to natural fiber cords were

evaluated by Fay and Lindquist (1954). They found sisal cord to be more attractive than jute or

wool cords of the same size, but less attractive than a similar sized cotton cord. When given a

choice between only cotton and sisal cords, flies preferred the sisal cord. They also found that

the same type of cord was more attractive to flies as the cord diameter increased between 0.13-

1.1 cm.

In our attractiveness experiment, the cords we evaluated varied by fiber type (animal or

plant), color, texture, and, in some cases, even diameter. All of the natural cords we evaluated

were more attractive than the synthetic cords. The natural cords were "rougher" than the

relatively smooth synthetic cords; in addition, the plant-fibered manila cord and the animal-

fibered leather cord were darker than the other cords. These factors may have increased their

overall attractiveness to the flies. If comparing the most attractive cord (manila) to the least

attractive (parachute cord) the differences in texture and color are substantial (Figure 3-4).

37









Manila is a very rough, coarsely textured brown thatch cord made from the leaf fibers of the

abaca tree, Musa textiles, while the parachute cord is a relatively smooth kemmantle cord made

of white nylon. The parachute cord was one of two cords less than 0.64 cm, which may have

decreased its attractiveness. The other cord less than 0.64 was the leather cord, but it was as

attractive as the other natural cords (except manila) despite its diameter being half the size. The

leather cord's dark color may have increased its attractiveness or it may have been more

attractive due to animal odors that were still associated with the material.

The previously available commercial fly cords were exclusively made of cotton. Cotton

was cheap, durable, absorbent and widely available. Although cotton was relatively attractive in

our experiments, it had very poor efficacy for both fipronil and indoxacarb when compared to the

other natural and synthetic cords tested indicating that it may not be the best type of cord to use

for insecticide treatment. Fipronil- and indoxacarb-impregnated wool cords had the greatest

efficacy in our experiments despite flies resting on it 50% less than the manila cord in the cord

attractiveness experiments. This is contrary to the previous assumption that quicker mortality

would result from increased exposure to a more attractive insecticide-impregnated cord and

neglects to account for the insecticide-substrate interaction. Highly organic materials readily

bind to pesticides and make them less effective (Dell et al. 1994, Gardner et al. 2000) and may

have accounted for the low LT50 and LT90 values seen in the cotton cords and in the LT50 value

of the fipronil-impregnated manila cord. The exact reason wool outperformed the other cords in

our experiments was not fully investigated, but it is likely due to the insecticide-substrate

interaction. The wool cord was the only animal-fibered cord evaluated and is naturally

impregnated with several oils. Both fipronil and indoxacarb are very lipophilic insecticides and









probably dissolved readily within these oils which likely increased the rate of insecticide transfer

from the cords through the waxy layers of the fly's cuticle.

In the laboratory, the indoxacarb cords generally provided a much slower kill than the

fipronil cords, however, in the field cages differences were not as apparent. Indoxacarb is a pro-

insecticide that needs to be bioactivated within the insect before it is toxic and will always cause

mortality slower than an insecticide, such as fipronil, that is toxic upon contact once a lethal dose

is obtained. Flies poisoned by indoxacarb in the laboratory are shielded from desiccation and

predation, which may have proved to be vital to their prolonged survival in the laboratory

experiments. Furthermore, the indoxacarb dose was increased in the field cage experiments and

may have affected the faster results seen in the field cage experiments. Both treatments showed

a decrease in efficacy in the field experiments after being aged 4 weeks, but still had adequate fly

count reductions and causing significantly more flies to die than the control.

In conclusion, the use of insecticide-impregnated cords is very practical to supplement a

house fly management program. Insecticide-impregnated cords ensure adequate residual

coverage in areas difficult to treat with traditional residual insecticides and they can easily be

removed and relocated to other fly resting areas if needed or alternated with cords impregnated

with other active ingredients to reduce the possibility of resistance development. More research

still needs to be done to determine adequate doses and rates of treatment, keeping in mind that

these may vary depending on cord type and insecticide used. Wool cord outperformed all other

cords evaluated in this study and fipronil and indoxacarb both appear to be effective insecticides

for house fly control.











Table 3-1. Efficacy of various cords impregnated with 0.1% fipronil or 0.6% indoxacarb on female house flies.


Lethal Times (h) (95% CL)


Treatment Cord
Fipronil Cotton
Manila
Wool
Polypro
Nylon


Potency Ratio (95% CL)


nt Slope SE


2750
1000
1250
2500
1500


9.52 + 0.36b
12.98 + 0.83a
5.32 + 0.29c
4.65 + 0.29d
3.68 + 0.25e


39.7 (39.2-40.2)e
35.0 (34.5-35.6)d
12.9 (12.3-13.4)a
26.2 (25.6-27.0)c
23.0 (21.2-24.6)b


54.1
44.0
22.4
49.6
51.3


(52.9 55.5)c
(42.6 45.7)b
(21.3- 23.8)a
(46.4- 53.8)c
(48.2 55.4)c


9.060
1.213
0.650
3.643
1.455


0.1067
0.5445
0.4200
0.7249
0.6927


1.00 e
1.13 (1.12-1.15)d
3.09 (3.01-3.16)a
1.51 (1.45-1.57)c
1.72 (1.66-1.79)b


1.00 d
1.23 (1.21-1.25)b
2.42 (2.35-2.48)a
1.09 (1.04-1.14)c
1.05 (1.00-1.11)cd


Indoxacarb Cotton 2248 4.04 + 0.13c 52.2 (50.3-54.3)c 108.5 (102.1-115.9)c 1.3494 0.5093 1.00 c 1.00 c
Manila 1659 5.10 + 0.74b 36.2 (32.3-38.7)ab 64.7 (60.2 73.4)b 3.4030 0.3336 1.44 (1.35-1.54)ab 1.68 (1.54-1.82)b
Wool 4250 6.44 + 0.16a 32.6 (32.0-33.1)a 51.5 (50.1 53.1)a 8.6295 0.2804 1.60 (1.54-1.67)a 2.11 (2.01-2.20)a
Polypro 2250 3.11 0.20d 52.2 (49.7-54.5)c 134.8 (122.7-151.7)d 0.6717 0.7147 1.00 (0.91-1.10)c 0.80 (0.72-0.90)c
Nylon 2000 6.57 0.54a 39.2 (37.2-40.8)b 61.5 (59.4 64.4)b 2.9321 0.2308 1.33 (1.27-1.39)b 1.76 (1.68-1.85)b
tTotal number of trials; 500 flies/trial except for the cotton (498) and manila (487) indoxacarb-impregnated cords (Probit [SAS Institute 2002]).
Mortality was corrected using Abbott's Formula. Means within a column, in the same treatment group, followed by the same letter are not significantly
different based on non-overlap of 95% confidence intervals.










Table 3-2. Cumulative number of dead flies and percent fly count reduction in relation to control fly counts of house flies exposed to
0.1% fipronil- and 1.2% indoxacarb-impregnated cords in field cages.
% Fly Count Reduction SEMI # of Dead Fliest
Treatment Age Treatment 24 h 48 h 24 h 48 h
0 Weeks Fipronil 80.22 13.10a 98.66 + 1.34a 83.0 + 1.0a 95.3 + 3.4a
Indoxacarb 57.39 + 6.92a 97.21 + 1.43a 30.3 4.4b 59.7 + 2.0b
Control 4.7 4.2c 11.3 8.5c

4 Weeks Fipronil 59.25 + 22.10a 87.43 12.57a 53.0 + 7.8a 79.3 7.2a
Indoxacarb 64.39 5.89a 87.72 7.34a 30.3 4.7b 62.0 + 8.1a
Control 3.3 + 1.7c 10.3 + 2.0b
t Cumulative mean number of flies recovered from cage floor.
Means in a column, within the same treatment age, followed by the same letter are not significantly different (P > 0.05; Student's T
or Student-Newman-Keuls test)






































I ss I


Laboratory and field experimental design elements. A). Laboratory arena
constructed of PVC pipe. Cords were suspended between the rubber bands using
paper clips. B) Sampling stage used in field cage experiments. Chick feeders with
either 10% sugar water or tap water and a plastic cup containing previously used
larval medium was used as sustenance and attraction. C). Cord placement in
relation to sampling stage (SS) in the field cage bioassay. Crossed circles were
short cords (0.6 m) and empty circles were long cords (0.9 m).


Figure 3-1.





























1 I I I


It


.44
V
*1~
VC~


4.
4 <


Figure 3-2. Attraction of female house flies to various natural and synthetic cords.


3.5


S5-
-. 3

. 2.5 -

S2 -
o
S1.5-

S 1-

0.5


It


; ;C-0


C

Sm-I


1 C


'IF








100 A
-i-]-

80

60- B
0 T
2 40 -

20 C

0 4 ---I1.




A.
100
A
T+ B
80- AB T

60- c
0
40o

20-
D




B. #

Figure 3-3. Female house fly mortality exposed to various natural and synthetic cords treated
with 0.1% fipronil for 24 h (A) and 0.6% indoxacarb for 48 h (B).














si'


Figure 3-4. Comparison of the most attractive cord (manila) and the least attractive cord (nylon
parachute) in the cord attractiveness experiments.









CHAPTER 4
EVALUATION OF A NEW IMIDACLOPRID BAIT FOR HOUSE FLY CONTROL

Introduction

The house fly, Musca domestic L., is the most commonly encountered pest of the

generalized group of Diptera called filth flies. Large numbers of house flies are frequently found

in areas where manure, garbage, and other decaying organic matter are abundant. Although

primarily a nuisance to people and animals, house flies can pose a health risk by mechanically

transferring pathogens picked up from their breeding sites, particularly when they enter homes or

eating establishments. When the source of infestation is inaccessible or sanitation measures are

not effective, house fly control is often achieved using dry insecticidal scatter baits.

Two widely used scatter baits are Maxforce Granular fly bait and Golden Malrin fly bait.

Maxforce Granular is an imidacloprid-based bait containing the fly attractant (Z)-9-tricozene,

the bittering agent Bitrex, and other attractants and inert ingredients. It is currently the only

imidacloprid-based scatter bait available. Golden Malrin contains 1.1 % methomyl, 0.049 %

(Z)-9-tricosene, as well as other attractants and inert ingredients, and is one of several methomyl-

based scatter bait formulations available.

In general, dry scatter baits have many advantages over other types of insecticidal fly

control products: they are easier to work with in field environments, they can be more attractive

to flies than liquid baits, and they usually have a longer storage shelf life (Gahan et al. 1954,

Darbro and Mullens 2004). However, dry scatter baits need to be replaced frequently in some

areas when granules become covered by manure or other debris (Barson 1987). The U.S.

Environmental Protection Agency (EPA), acting under legislative mandates, also requires the

scatter bait granules be dyed to distinguish them from other non-toxic materials. For example,

the Maxforce Granular fly bait is formulated as red granules and the Golden Malrin is









formulated as blue granules. When these granules become wet, the dye often bleeds onto the

surrounding surface and may be unsightly for the user.

Label restrictions are also very different and can limit the uses of certain active ingredients

or insecticide products. Golden Malrin can only be applied as scatter bait or within bait

stations, whereas, the Maxforce Granular can be applied as scatter bait, within bait stations, or it

can be mixed with water and painted onto surfaces allowing it to be applied directly to distinct

fly resting areas, such as on ceilings or rafters. However the use of Maxforce Granular is more

restricted than that of Golden Malrin because its label restricts its use in food establishments.

Golden Malrin, despite being the only carbamate-based insecticide not classified as "restricted-

use", can be used within food establishments when used in bait stations placed at least 1.2 m

from the ground in areas where food processing or preparation does not occur.

An imidacloprid sprayable bait, Maxforce Fly Spot, has recently become commercially

available. It contains 10% imidacloprid, 0.1% Z-9-tricosene, Bitrex, and inert ingredients. This

formulation still maintains the advantages of traditional scatter baits, while eliminating some of

the disadvantages of the currently available products. Once applied, Maxforce Fly Spot bait

dries clear and the label allows for application within food establishments when the facility is not

in operation.

Our objectives were to compare the effectiveness of the new sprayable bait in relation to

the two most commonly used dry scatter baits. In addition, we compared the performance of the

imidacloprid sprayable and granular baits in a controlled field environment and tested the

imidacloprid sprayable bait impregnated in cords.









Materials and Methods

Insects. The Horse Teaching Unit (HTU) strain of house flies, M. domestic L., reared at

the University of Florida in Gainesville was used for all experiments. Larvae were reared on a

diet medium, modified from Hogsette (1992), containing 3 liters wheat bran, 15 ml methyl

paraben, 1.5 liters water, and approximately 200 g (250 ml) dairy calf feed (Calf Manna

pellets, Manna Pro Corp., St. Louis, MO). All developmental stages were held at 26 + 1IC and

55% RH with a 12:12 (L:D) photoperiod. Adult flies emerged within in screened rearing cages

and were provided granulated sugar, powdered milk, and water ad libitum.

For all assays, adult house flies (3-5 d old) were aspirated from the screened rearing cages

using a handheld vacuum with a modified crevice tool attachment. Flies used for the laboratory

assays were placed into a 50C environment for 5 min to subdue activity. Flies were then placed

on a chilled aluminum tray, sexed, and counted. House flies used for field cage assays were not

subdued, but were aspirated from the screened rearing cages and released directly into field

cages.

Laboratory Arena Design. Arenas (31 x 25 x 21 cm) were constructed using PVC pipe

(1.27 cm [1/2 in] diam) (Figure 3-1A). Rubber bands were used to establish five uniformly

distributed cord positions along the length of the arena. Each position held one cord which was

vertically attached to the rubber bands using paper clips. Five cords were used with all

laboratory experiments: nylon (Braided, Multi-Purpose Braid 75 lb. load limit, Wellington

Cordage LLC, Madison, GA), polypropylene (Braided, Multi-Purpose Rope 56 lb. load limit,

Wellington Cordage LLC, Madison, GA), cotton (Braided, Multi-Purpose Sash Cord 28 lb.

load limit, Wellington Cordage LLC, Madison, GA), manila (Twisted, Natural Rope 108 lb.

load limit, Wellington Cordage LLC, Madison, GA), and wool (Twisted, Natural Cord, Wooded









Hamlet Designs, Greencastle, PA). Arenas were enclosed with a transparent plastic bag (3716

cm2 [24 x 24 in], 1 mil poly, Uline, Waukegan, IL).

Field Cage Design. Cages (1.8 x 3.7 x 1.8 m) were constructed from PVC pipe (2.54 cm

[1 in] diam) and enclosed with mesh screening (Outdoor Cage, #1412A, 18 x 14 mesh, Bioquip,

Rancho Dominguez, CA). Black plastic sheeting (6 mil) was used to line the floor. A sampling

stage, constructed of two vertical cinder blocks and an inverted storage bin (Palletote #1721, 37

liter, Rubbermaid, Winchester, VA), was placed in the center of the cage (Figure 3-1B). On top

of the sampling stage there were two 994-ml (1 qt) chick waterers, one filled with 10% sugar

water and the other with tap water, and a 60-ml plastic cup filled with 8 g of previously used

larval house fly medium. The chick waterers provided enough sustenance for the duration of the

test and the plastic cup was used as an attractant. The plastic cup was covered with a paper towel

and sealed with a rubber band to prevent flies from ovipositing on the medium.

Fly Bait Comparisons. Three fly baits, 2 dry scatter baits and 1 sprayable bait, were

applied to polystyrene Petri dishes (100 by 15 mm; Fisher Scientific, Pittsburgh, PA). The

methomyl granular bait (Golden Malrin, Methomyl 1.1%, (Z)-9-Tricosene 0.049%, Wellmark

International, Schaumburg, Illinois; dose: 0.23 g/0.9 m2) and the imidacloprid granular bait

(Maxforce Granular fly bait, Bayer CropScience, Kansas City, MO; dose: 30.17 g/0.9 m2)

were sprinkled on the Petri dish. The Imidacloprid sprayable bait (Maxforce Fly Spot bait,

Imidacloprid WG 10, Lab Code: 342/207-7, Bayer CropScience, Monheim am Rhein, Germany;

dose: 0.45 g/0.9 m2; rate: 0.12 g Pr/ml/0.9 m2) was suspended in tap water, sprayed on the Petri

dish bottom using an airbrush (Paasche, Type H, Chicago, IL), and allowed to dry in a fume

hood prior to being placed in the arena. Bait dishes were placed on the bottom rubber bands in

the center of the arena. A separate arena with an untreated Petri dish was used as the control.









Cords in these experiments served only as resting positions for the flies, they were untreated and

hung in the same configuration for all repetitions: (left to right: position 1 = cotton; 2 = wool; 3

manila; 4 = polypropylene; 5 = nylon).

Groups of 50 female flies were placed within each arena and a 10% sugar water solution

was provided ad libitum. Mortality was recorded at 1, 3, 5, and 24 h. Flies were considered

dead if they were unable to stand or fly. Each experiment was run in the laboratory (30 1C)

under continuous light and replicated three times.

The two imidacloprid baits were evaluated in the field cages. Treatments consisted of two

plastic lattice squares (0.19 m2) treated with imidacloprid granular bait, imidacloprid sprayable

bait, or tap water (control). Treatments were applied on only one side of the lattice at the same

rates as the laboratory fly bait comparison assays. The imidacloprid granular bait was mixed

with tap water (1.44 g: 1 ml) and painted on. The tap water (3.78 ml/0.9 m2) and imidacloprid

sprayable bait (0.12 g Pr/ml/0.9 m2) were sprayed on using an airbrush. All treatments were

allowed to thoroughly dry outdoors in the open air before being hung on the ceiling PVC pipes

using cable ties. Each lattice square was placed medially along the length of the cage,

approximately 0.5 m away from each side of the sampling stage and positioned so that the

treated surfaces of the lattice squares faced opposite directions.

Depending on fly availability, 27.5 35 ml (9.8 + 1.8 flies/ml) of flies were released into

each cage. After a 1-h acclimation period, pre-treatment fly counts were taken. Before fly

counts were taken, the operator walked three laps around the interior of the cage to disturb flies

from their resting positions and to recover any dead flies from the cage floor. Four consecutive

fly counts were then taken from the outside of the cage 1 min after exiting. All flies that landed

on the sampling stage, chick waterers, and plastic cup attractant were counted. Treatments were









then hung within the cages and post-treatment fly counts were taken at 1 and 24 h using the same

method described above. After the initial 24 h evaluation, treatments were aged in the elements

for two weeks, at which point residual effectiveness was re-evaluated as described above. Three

replicates were performed at each treatment age (0 and 2 wk).

Bait-Treated Cords. Five laboratory arenas were organized into a 5 x 5 Latin square

design, blocking for treatment position, and a sixth arena was used as the control. Each

treatment consisted of a cord (15.2 cm length, 0.6 cm diam) impregnated with a 2.5% solution of

imidacloprid sprayable bait. The imidacloprid solution was prepared by combining 25 g of the

formulated insecticide with 100 ml of tap water. Cords were impregnated by dipping for -2 sec

in the insecticide solution and were then allowed to dry on aluminum foil covered trays in a fume

hood prior to being placed into the arenas. The control arena had no treated cords and had the

same cord configuration as the fly bait comparison bioassay described above.

Laboratory tests were conducted with groups of 60 female flies/arena. Flies were provided

a 10% sugar water solution ad libitum. Morbidity (knockdown) was recorded at 2-5 h post-

treatment and mortality was recorded at 24, 48, and 72 h. Flies were considered knocked down

if they did not move when touched at the 2-5 h recordings. Flies that were unresponsive to touch

at the 24, 48, and 72 h recordings were considered dead. Each experiment was run under

continuous light in the same laboratory conditions as described above and replicated twice.

In the field cages, treatments consisted of two long (0.9 m) and eight short (0.6 m) lengths

of imidacloprid-impregnated wool cords. Each cord was treated in the same manner as the

laboratory experiments, except the cords were dipped and soaked for 1 min prior to drying. Flies

were released and fly counts were taken in the same manner as the imidacloprid bait field cage

experiments. Cords were hung vertically from the mesh ceiling using paper clips in specific









locations, which remained constant throughout the experiment (Figure 3-1C). Post-treatment

sampling counts were done at 24 and 48 hrs. After the initial 48 h evaluation, treatments were

aged in the elements for four weeks, at which point residual effectiveness was re-evaluated as

described above. Three replicates were performed for each treatment age (0 and 4 wk).

Data Analysis. All analyses were done using a one-way analysis of variance with JMP

IN (SAS Institute 2005). For the fly bait comparison and the bait-treated cord experiments,

percent morbidity (bait-treated cords) and mortality data were arcsine square root-transformed

and analyzed for each time interval. For the field cage experiments, percent fly count reductions

were calculated from the control fly counts. Fly count reductions and mortality data (number of

dead flies recovered from cage floor) were then analyzed for each treatment age (0 and 2 wk for

the imidacloprid comparisons; 0 and 4 wk for the bait-treated cord experiments) (Conover and

Iman 1981). Means for all analyses were separated using the Student's T test or the Student

Newman Kuels (SNK) method (a = 0.05).

Results

In all laboratory experiments, flies did not fully recover from chilling until roughly 1 h

after entry into the arenas. Flies first contacted the baited Petri dishes in the bait comparison

experiments approximately 35 min post recovery in the following order: imidacloprid granular

bait, imidacloprid sprayable bait, methomyl granular bait. Initial fly contact on the treated cords

in the bait-treated cord experiments was not observed.

Fly Bait Comparisons. The imidacloprid granular and the imidacloprid sprayable baits

had higher fly mortality than the methomyl granular fly bait at 3 h, but by 24 h the methomyl

granular bait had the highest overall mortality (Figure 4-1). At 24 h, fly mortality with the

imidacloprid sprayable bait was not significantly different from mortality with either the

imidacloprid granular or the methomyl granular fly baits, but significant difference in fly









mortality did exist between the methomyl and the imidacloprid granular fly baits. All treatments

were significantly different than the control fly mortality at all observations > 3 h.

In the imidacloprid field cage experiments, no differences were seen between either

treatments with fresh or aged cords (Table 4-1). Both treatments had >35% fly count reductions

at 24 h and >70% fly count reductions by 48 h with fresh cords, but fly count reductions did not

exceed 8% with aged cords for either treatment. A fly count increase was observed with the

imidacloprid granular treatment at 48 h with aged cords. The number of dead flies collected in

the treatment cages was significantly different from the control with fresh cords, but no

differences were observed between the aged treatments and controls 2 wk post-treatment.

Bait-Treated Cords. Morbidity increased on a time-dependent basis until approximately

3-4 hours post-treatment, at which time flies recovered from being knocked down by all of the

imidacloprid bait-treated cords except for the cotton cord (Figure 4-2). Flies exposed to all

imidacloprid bait-treated cords had knockdown recovery by 24 h. All cords caused significantly

more mortality than the control cords at every 24 h recording (Figure 4-3). Beyond 24 h,

mortality with the imidacloprid bait-treated nylon, cotton, and wool cords increased more sharply

than the mortality caused by the bait-treated manila and polypropylene cords. The imidacloprid

bait-treated wool cord caused the highest overall fly mortality (74%). All other cords resulted in

house fly mortalities <60% with the imidacloprid bait-treated polypropylene cord showing the

lowest overall fly mortality (25%).

In the field cages, the imidacloprid bait-treated cords caused >87% fly count reductions by

24 h with fresh and aged cords (Table 4-2). The aged bait-treated cords fly count reductions

decreased by -6% by 48 h, whereas fly count reductions increased by -6% with the fresh cords.









The number of dead flies collected was significantly different than the control at 24 and 48 h for

both fresh and aged cords, except for the 48 h recording with the aged cords.

Discussion

When insecticides are used for house fly control, most users expect to see satisfactory

results (i.e. dead flies) within hours and markedly reduced populations within 1-2 days. Thus, an

effective fly bait will attract flies quickly and cause high mortality within a relatively short

period of time. In our bait comparison experiment, flies contacted the imidacloprid baits sooner

than they contacted the methomyl bait, which may have been a contributing factor to the higher

fly mortality at 3 h with the imidacloprid baits than with the methomyl bait. However, the higher

fly mortality with the methomyl bait after 24 h suggests that methomyl may be a more potent,

although slower acting, active ingredient.

Other studies comparing imidacloprid and methomyl baits have also shown the same

mortality trends we observed in flies exposed to technical and bait formulations of imidacloprid

and methomyl (White et al. 2007). In those experiments, White et al. observed up to 50% of the

flies that were knocked down by imidacloprid formulations recovered. They hypothesized innate

characteristics, independent of resistance mechanisms, may make some flies tolerant to

neonicotinoids. We observed knockdown recovery in the bait-treated cord experiments with all

cords, but no recovery was seen in the house flies exposed to any of the baits we tested in the

bait comparison experiments. Recovery may have occurred in these experiments, but was not

observed because recordings were not taken between the 5 h and 24 h recordings. Flies that were

knocked down were not isolated from the arena in our experiments and could have received a

second dosing before having the opportunity to fully recover.

Differences in cord material or treatment application technique may have also attributed to

fly recovery in the bait-treated cord experiments. Cord saturation is dependent on the cord









composition and may have lead to sublethal dosing. We observed that cord composition varied

between the types of cords we used, and even among individual cords. Distribution of oils and

other materials on the surface of each cord make it difficult to have the bait uniformly distributed

over the surface of the cord. When bait is sprayed onto a solid surface, such as the Petri dish, a

precise amount of bait remains on the surface after the water evaporates. However, when a cord

is dipped into a bait solution, the bait may be absorbed deep into the fibers, disperse throughout,

or pool in areas on the cord. Thus, some bait may not be available for flies to contact. This is

evident when hand-dipping cords in dyed insecticides materials, such as in an indoxacarb

wettable granule (WG) solution, which is grayish-brown in color. Despite being fully

submerged in the insecticide solutions, some of the cord often remains its natural color,

apparently void of any bait. Once the same cords are allowed to completely dry and are removed

from the drying trays, brown staining surrounds were they once lay indicating that some of the

insecticide may be lost during the drying process as well.

It is undetermined if knockdown recovery occurred in our field cage experiments. If flies

are knocked down in the field, natural enemies may prey upon them before they are able to fully

recover. We observed knocked-down flies being preyed upon by ants, spiders, and lizards.

Others, undoubtedly, became victim to desiccation after being knocked down. Barson (1987)

found that flies knocked down by methomyl in the field often lost their ability to fly, but still had

the ability to reproduce. White et al. (2007) commented that 10% of the flies knocked-down by

imidacloprid in their laboratory studies fully recovered and resumed normal behavior when

protected from a second exposure to imidacloprid. The inability to fly would make flies more

vulnerable to predation by natural enemies. However, if reproduction is still occurring, it will be

detrimental to any fly control program because of a fly's prolific reproductive capabilities. Field









studies to determine the effect of knock down and recovery on a fly management program would

be beneficial.

Insecticide-impregnated cords have been used extensively in the past to control flies and

have recently been examined using new insecticides not yet registered for use against house flies

(Unpublished, Chapter 3). In those experiments, indoxacarb and fipronil were more effective on

wool cords than any other natural and synthetic cords tested. Wool cords also showed higher

efficacy than the other natural and synthetic cords tested when treated with the imidacloprid

sprayable bait. Exact LT50's were not determined in these experiments because of the

knockdown and subsequent recovery observed, but based on Figure 4-3, we estimate that 50% of

the flies died after approximately 60 hours (2.5 d) with the wool cord. With such a long period

to reach 50% fly mortality, imidacloprid-treated cords were slower acting than any of the

fipronil- or indoxacarb-impregnated cords previously tested. However, in the field cages, the

imidacloprid bait-treated cords reduced fly counts by 80% in 24 h. The high lipid content of

wool cord may facilitate the transfer of insecticide through the cuticle of the house fly, but this

does not explain the differences in results between the laboratory and field assessments.

Differences in the rate of cords per cage area may explain the differences in the laboratory and

field results. The cords in the field cages were hung at a rate of 9.1 m of cord/9.3 m2 area based

on the recommended rate of the insecticide cords used in the 1950's (Fehn 1958, Smith 1958,

Weinburgh et al. 1961). The rate in the laboratory arena was comparatively much lower, 0.02 m

of cord/m2 vs. 1.0 m of cord/m2. This rate appears to be quite high and may vary between

different cord/insecticide combinations. Additionally, the aforementioned predation and

desiccation of the knocked down flies probably was significant factor contributing to the rapid

fly count reductions in the field cages.









When comparing the imidacloprid bait-treated cords and the imidacloprid bait-treated

lattice squares, the bait-treated cords were more effective. The lattice squares did not reduce the

fly counts to any significant degree after being aged 2 weeks, but the imidacloprid bait-treated

wool cords had good fly count reductions even after being aged 4 weeks. The plastic lattice

squares were selected as a treatment surface to represent the material found on many portable

toilets, latrines, or dumpsters. Bait treatments on this type of surface are very vulnerable to

environmental conditions because the material does not allow the bait to penetrate as in the cord

treatments. Damp conditions in the mornings and unexpected precipitation (6.5 cm) that

occurred between evaluation intervals washed away most of the bait product from the lattice

squares. With the imidacloprid granular bait application, the lattice squares were almost

completely void of the red dye following these moisture events and red staining was seen on the

cage floor. We assume that the imidacloprid sprayable bait was also washed off the lattice

squares given the results of the fly count reductions and the dead flies recovered, but was not

observed because the bait has no color. The bait-treated cords were exposed to 3.5 cm less

precipitation than the bait-treated lattice squares, which may have also affected the bait available

on the cord. When dipping cords in a bait solution, the bait is absorbed in between individual

cord fibers and even deeper into the core of the cord, making the bait more protected from

environmental conditions. When the cords are then subjected to these moisture events, the bait

may concentrate in specific areas of the cord (such as the cord end) instead of completely leaving

the cord as seen with the lattice. Additionally, flies prefer to rest on cords and probably receive a

larger dose of insecticide in this manner as compared to when they land on the flat surfaces of

the lattice. When a fly lands on a flat surface they are exposed only to the precise amount of









toxicant that absorbs through their tarsi or is imbibed; however, when resting on cords, their

thorax and abdomen are also brushed by the treated cord fibers.

In conclusion, the imidacloprid sprayable bait was found to be as effective as the

traditional commercial scatterbaits compared in this study. Its unique formulation and less

restrictive product label allow this bait to be used in areas where other fly baits are prohibited.

Unless a more rain-fast formulation becomes available, the imidacloprid sprayable bait will need

to be reapplied frequently in areas with high moisture or precipitation especially when applied to

non-absorbent surfaces such as portable latrines or dumpster lids. The bait's potential

effectiveness in insecticide-impregnated cords needs further investigation due to differing

laboratory and field results. Regardless, this new imidacloprid sprayable bait should prove to be

a very useful tool in any fly management program.









Table 4-1. Number of dead and percent fly count reduction in relation to control fly counts of house flies exposed to imidacloprid
bait-treated lattice squares in field cages.


Treatment Age
0 Weeks


Treatment
Imidacloprid granular bait
Imidacloprid sprayable bait
Control


% Fly Count Reduction SEMI
1 h 24 h
47.1 + 6.3a 70.9 + 4.4a
36.6 + 20.5a 80.2 + 4.7a


# of Dead Fliest
lh
36.0 10.0a
36.3 2.0a
0.3 + 0.3b


24 h
117.0+ 9.5a
113.0 10.la
1.7 0.7b


2 Weeks Imidacloprid granular bait 0.8 8.2a -3.8 20.1a 1.0 0.6a 19.7 11.2a
Imidacloprid sprayable bait 7.6 10.8a 6.3 + 19.5a 0.7 + 0.7a 14.0 + 11.6a
t Mean number of individuals recovered from cage floor.
Means in a column, within the same treatment age, followed by the same letter are not significantly different (P > 0.05; Student's
T test or Student-Newman Kuels Method)












Table 4-2. Number of dead and percent fly count reduction in relation to control fly counts of house flies exposed to
imidacloprid bait-treated cords in field cages.
% Fly Count Reduction SEM # of Dead Fliest
Treatment Age Treatment 24 h 48 h 24 h 48 h


0 Weeks Bait-Treated Cords
Control

4 Weeks Bait-Treated Cords
Control


90.4 + 4.1


87.9 + 0.9


96.8 3.2


82.4 14.7


97.7 17.2a
12.7 + 4.6b

50.7 14.0a
8.7 3.8b


114.3 19.8a
19.7 7.7b

69.3 + 23.7a
13.0 4.7a


t Mean number of individuals recovered from cage floor. Means in a column, within the same treatment age, followed by the
same letter are not significantly different (P > 0.05; Student's T test)
















- 4- Methomyl Granular
- -- Imidacloprid Granular
--Imidacloprid Spray able
-A- Control


.. S -


-


0 5 10 15 20 25
Time (h)



Figure 4-1. Mortality of female house flies exposed to imidacloprid and methomyl granular
scatter baits and a sprayable imidacloprid bait.


0











20
_-m Cotton (plant)
18 -0 Manila (plant)
Wool (animal)
16 -A Nylon (synthetic)
Polypropylene (synthetic)
14 + Control

S12

S10-
8
-7-

6 f

4


0 1 2 3 4
Time (h)


Figure 4-2.


Morbidity (knockdown) of female house flies exposed to natural and synthetic
cords dipped in a 2.5% solution of imidacloprid sprayable bait.













90
80
70
S60
S 50
40
30
20
10
10 -
0
0


Figure 4-3.


-*- Cotton (plant)
-D Manila (plant)
- Wool (animal)
- -k Nylon (synthetic)
-A- Polypropylene (synthetic)
- Control


24 Time (h) 48
Time (h)


Mortality of female house flies exposed to natural and synthetic cords dipped in a
2.5% solution of imidacloprid sprayable bait.









CHAPTER 5
SUMMARY AND CONCLUSIONS

House flies are often found in tremendous numbers in situations where U.S. troops are

most frequently deployed, such as areas stressed by conflict or natural disasters. House flies are

nuisance pests that decreases troop morale and they pose a health risk to deployed troops that

may disrupt mission objectives if diseases that are associated with their presence stress the health

care systems. The main objective for this research was to evaluate insecticide-impregnated cords

and sprayable fly bait as new methods to control the house fly and provide usable information to

the DOD that will help protect the deployed war fighter.

First and foremost, house flies can be controlled using insecticide-impregnated cords and

sprayable fly bait and their use would benefit agricultural, urban, and military fly management

programs. Insecticide-impregnated cords and sprayable fly baits are both very easy to use

products. Impregnated cords can be hung using a staple gun or other similar method and

sprayable bait is mixed with water and sprayed onto any surface. Impregnated cords can be

removed and relocated quickly, which may be beneficial in relatively mobile troop deployments

or in situations were resistance is suspected. Sprayable baits can be removed with simple water

wash down and easily reapplied when needed.

One of the most interesting findings in this research is the further understanding in

insecticide-impregnated cord toxicity. Previous insecticide-impregnated cords were made

exclusively of cotton because it was cheap, durable, and relatively attractive to house flies,

however, the cotton cords in our experiments were the least effective cord. Wool cords

consistently showed they were more effective than any of the other cords we evaluated despite

the fact that they were not the most attractive cord to the house flies. House flies were most

attracted to the highly organic manila cord and preferred to rest on it more than any other natural









or synthetic cord evaluated. The wool cord was the only animal-based cord impregnated with

insecticides and possibly gave it a distinct advantage over the other cords because it is naturally

coated with oil, which probably helped facilitate insecticide transfer through the fly cuticle.

Two fly scatter baits, Maxforce Granular and Golden Malrin fly bait, are both listed on

the DOD pesticide contingency list for house fly control but both have limitations. Only one can

be applied in food service areas and both can stain materials and equipment which can

compromise camouflage and substrate appearance. Eliminating these disadvantages, while

maintaining bait efficacy, would appear to provide advantages to a fly management program and

benefit the deployed war fighter. The new sprayable imidacloprid bait, Maxforce Fly Spot, is

as effective as the two previously mentioned scatter baits in the laboratory and as effective as its

counterpart, Maxforce Granular, in the field. Its unique formulation allows it to be used within

food serving areas and it will go unnoticed because it dries clear. Unfortunately, like the other

bait products, environmental factors, such as rain, decrease the efficacy over time. Residual

efficacy did improve when the bait was applied to the cords rather than the non-absorbent plastic

surfaces often found on latrines and dumpsters.

We anticipated that the research completed here would provide some information that

could be further used to develop future products that could be benefit the DOD. The insecticides

evaluated in the impregnated cord studies are both non-registered for house fly control. House

flies have shown little to no resistance towards fipronil and indoxacarb and both insecticides

appear to be very effective against this pest. At this time, no information has been obtained on

whether or not the manufacturers of fipronil or indoxacarb are seeking registrations for these

products to control flies. However, a Colorado company has informed me of their interest in

indoxacarb-impregnated cords and has begun conversations with DuPont regarding further









research into this type of product. The sprayable imidacloprid fly bait is not currently listed on

the DOD pesticide contingency list but it received its EPA registration in 2006 and became

commercially available in mid 2007. A formal request will be submitted to the Armed Forces

Pest Management Board this summer to request that it be assigned a National Stock Number

(NSN) and be placed on the DOD pesticide contingency list. If successful, Maxforce Fly Spot

will be readily available to the deployed war fighter for use in their fly management programs.









APPENDIX A
REVIEW OF INSECTICIDE-IMPREGNATED CORDS

The use of insecticide-impregnated cords to control house flies was first tried with DDT in

1947 (Baker et al.). By the early 1950's, impregnated materials for fly control became

increasingly common (Pimentel et al. 1951). Insecticide-impregnated cords were being used on

dairies, at rural residences, military mess halls, state fairs, and state prisons with great success

(Kilpatrick 1955, Maier and Mathis 1955, Soroker 1955, Kilpatrick and Schoof 1956).

Commercial Fly-Cords distributed by Fly-Cord Inc. (Savannah, Georgia) were widely available

and used by 1957 (Fehn 1958, Smith 1958). These cords were considered the treatment of

choice for use in buildings housing animals because of the economy and efficiency (Fay and

Kilpatrick 1958).

Fay and Lindquist (1954) recognized that impregnated cords offer only a small percentage

of the surfaces available for flies to rest. Exploiting the factors which enhance a fly's attraction

to a particular cord would subsequently lead to higher mortality on impregnated cords. They

found that cord type, thickness, and color significantly influenced a fly's attraction to a particular

cord. Sisal and cotton cords were more attractive than jute or wool cords. Cord attractiveness

increased with cord diameters between 3/64" and 7/16". Flies preferred red and black cords over

blue, yellow, green, or white cords. No preference was evident between vertically or

horizontally hung cords.

Although several other organophosphate insecticides have been evaluated for their

effectiveness, all provided satisfactory results. However, only one cord was available

commercially (Kilpatrick and Schoof 1959, Gratz et al. 1964, Rabari and Patel 1976). The

commercial Fly-Cord was a 3/32" diameter, red cotton cord impregnated with 13.79% parathion

and 3.54% diazinon (Fehn 1958, Smith 1958). The cord was supplied on a reel containing 300









feet; each linear foot of cord contained 75-100 mg of parathion (Youngblood 1960). The

manufacturer recommended a rate of 30 linear feet of Fly-Cord per 100 square feet of floor area

in locations were adult flies congregate (Fehn 1958, Smith 1958). Cords were normally hung

about three feet apart using staple guns or simply tied on to the structure (Fehn 1958,

Youngblood 1960). If multiple competing resting surfaces were available to the flies, most

applicators increased the amount of impregnated cords in the area. In places where flies were

not seen resting or where conditions were unsuitable (i.e. areas with drafts), no cords were

placed.

The use of impregnated-cords was an attempt to find new methods to reverse the resistance

associated with residual spraying of chlorinated hyrdrocarbons such as DDT (Keiding and

Jespersen 1986). Fly-Cords offered an easy-to-use control method that restricted and

concentrated a residual insecticide. This method lowered the selection pressure for resistance

because flies which avoided contact with the cords diluted the remaining population of resistant

individuals (Keiding and Jespersen 1986). Other methods, such as paint-on baits, non-residual

space sprays, and combining baits and larvicides, were evaluated and determined effective

(Keiding and Jespersen 1986). Paint-on baits and non-residual space sprays are widely available

and used today in the United States.

No commercially available insecticide-impregnated cord products are currently available

in the United States. This is partially because the main active ingredients, parathion and

diazinon, are not registered for filth fly control. In addition, the use of selective insecticidal baits

has become increasingly popular. In Denmark, the use of impregnated cords was abandoned

because of newer construction techniques that allowed for more ventilation in the animal shelters

and more effective residual sprays became available (Keiding and Jespersen 1986). The World









Health Organization and the United States Military continue to recommend the use of insecticide

treated or impregnated cords (Rozendaal 1997, AFPMB 2006)









APPENDIX B
REVIEW OF FLY BAITS

Insecticidal baits have long been used for fly control. One of the original bait formulations

contained either 1-2% formaldehyde or sodium arsenite mixed with milk or sugar water (Keiding

1976). Residual insecticides have been mixed with sugar to make them more attractive to flies

and serve as a type of bait, but this type of mixture is often not as effective as baits specifically

formulated to attract flies. Today's fly baits are loaded with many different attractants including

pheromones, sugars, and other substances that specifically attract house flies. Most of these fly

baits are formulated as either dry scatter baits, but many also come in easy-to-use bait station

devices. Some of the dry scatter baits can be mixed with water and painted on a surface.

Insecticidal baits have many advantages over other chemical control methods. Baits are

relatively inexpensive, usually have a longer storage shelf life, are more attractive to flies than

other chemical control methods, and are easier to work with in field environments (Gahan et al.

1954, Darbro and Mullens 2004). The dry scatter baits simply get scattered on the ground in the

infested areas or placed within a bait station while bait station devices normally only need to be

opened and hung in infested locations. Keiding (1976) considered baits less likely to select for

resistance than residual sprays. He was most likely referring to the physiological resistance seen

in many of the organophosphate and carbamate insecticides at that time. Today, many insects,

including flies, have been shown to develop behavioral resistance to baits (Darbro and Mullens

2004).

Fly baits do have disadvantages. Dry scatter baits do not target fly resting areas unless

they can be painted on and this type of application often leads to stained surfaces because the

U.S. Environmental Protection Agency (EPA) requires the scatter bait granules to be dyed to

distinguish them from other non-toxic materials. Staining can also occur when these granules









become wet by rain and bleed onto the surrounding surface, which may be considered unsightly

for the user. Bait station devices are also degrade rapidly in environments with intense sun or

precipitation. Dry scatter baits also need to be replaced frequently in some areas when granules

become covered by manure or other debris such as garbage (Barson 1987).

As with any chemical insecticide, label restrictions can be disadvantageous (although

necessary in most cases) and limit the needs of the applicator. For example, the two most widely

used dry scatter baits in the United States are Maxforce Granular fly bait and Golden Malrin

fly bait. Maxforce Granular is an imidacloprid-based bait (neonicotinoid class) containing the

fly attractant (Z)-9-tricozene, the bittering agent Bitrex, and other attractants and inert

ingredients. Golden Malrin contains 1.1 % methomyl carbamatee class), 0.049 % (Z)-9-

tricosene, as well as other attractants and inert ingredients, and is one of several methomyl-based

scatter baits available. Maxforce Granular has a more restricted label than that of Golden

Malrin because its label restricts its use in food establishments. Golden Malrin, despite being

the only carbamate-based insecticide not classified as a "restricted-use" insecticide, can be used

within food establishments when used in bait stations placed at least 1.2 m from the ground in

areas where food processing or preparation does not occur.

Recently, a new fly bait has become commercially available that may offer advantages

over some of the other current fly baits available. Maxforce Fly Spot bait contains 10%

imidacloprid, 0.1% Z-9-tricosene, Bitrex, and inert ingredients. Once applied, Maxforce Fly

Spot bait dries clear and the label allows for application within agricultural livestock production

facilities and serving areas of food establishments when the facility is not in operation.









APPENDIX C
REVIEW OF INSECTICIDES EVALUATED

Fipronil

Fipronil is a phenylpyrazole insecticide that was first registered in the United States in

1996 (Connelly 2001). It is used to control termites, ants, roaches, fleas, ticks, and various other

agriculture and turf pests. No fipronil products are currently registered for house fly control.

Fipronil causes mortality by contact and ingestion (Vargas et al. 2005). Insects exposed to

fipronil show extreme neural excitation that eventually leads to insect paralysis and death. Death

is caused by the disruption of the normal passage of chloride ions through the y-aminobutyric

acid type A (GABA) receptor system of insects (Scharf et al. 2000). Hainzl et al. showed

fipronil to have a tighter binding affinity toward insect GABA-regulated chloride channels over

mammalian receptors (1998). Fipronil-sulfone, an important active metabolite of fipronil, was

also found to block the glutamate receptors in cockroaches (Zhao et al. 2004). Glutamate-gated

chloride channels are only found in invertebrate systems at skeletal neuromuscular junctions of

both the peripheral and central nervous system (Raymond and Sattelle 2002, Scharf 2003). The

unique quality of fipronil to affect two target sites makes it a highly selective insecticide and

potentially important factor limiting the development of detectable resistance (Zhao et al. 2004).

To date, resistance to fipronil appears to remain at low levels or even be non-existent in

house flies (Scott and Wen 1997, Scott et al. 2000, Kristensen et al. 2004). Low levels of cross-

resistance have been reported in multi-resistant house flies and attributed to monoxygenase-

mediated detoxification, decreased insecticide penetration, and target site mutations (Wen and

Scott 1999, Liu and Yue 2000, Kristensen et al. 2004). Resistance surveys in New York found

house flies susceptible to fipronil even at LC99 levels (Scott et al. 2000).









Indoxacarb

Indoxacarb (DPX-MP062) is an oxdiazine insecticide that was first registered in the United

States in 2000 (EPA 2000). It is a 75:25 mixture of the active S-isomer (DPX-KN128) and the

inactive R-isomer (DPX-KN127). A less effective formulation (DPX-JW062) contains a 50:50

mixture of the two stereoisomers. Indoxacarb was originally formulated to control lepidopteran

pests of fruits and vegetables, but newer registrations include cockroach, mole cricket, and fire

ant baits. It is not currently registered for house flies.

Indoxacarb is considered an organophosphate replacement and designated as a "reduced-

risk" insecticide by the EPA. It is a pro-insecticide that must be biochemically converted to a

toxic decarbo-methoxyllated metabolite (Dias 2006). Toxicological effects are dependent on the

conversion of the inactive metabolite to its toxic form within the insect body. In mammals,

indoxacarb metabolites are rapidly excreted; whereas in insects, indoxacarb is rapidly converted

by an esterase and amidase into DCJW, which is the more insecticidally active metabolite.

Insects exposed to lethal doses of indoxacarb experience impaired nerve function, feeding

cessation, paralysis, and eventually death. Indoxacarb poisoning occurs through contact or

ingestion and it works by blocking the sodium channel of the insect nervous system. This mode

of action is distinct from other insecticides that target the sodium channels of insects (DDT,

pyrethroids) because DCJW disrupts the sodium channels without modifying the activation or

deactivation kinetics (Lapied et al. 2001). It works by blocking the channel pore, and prevents

normal sodium ion flow.

Because indoxacarb is a new chemistry, not much work has been done on insecticide

resistance. Shono et al. (2004) selected house flies that had >118-fold resistance in as little as

three generations and concluded that the resistance mechanism was associated with a major









factor on autosome 4 and a minor factor located on autosome 3, both of which are not linked to

any resistance mechanisms previously described.

Imidacloprid

Imidacloprid is a chloronicotinyl nitroguanidine (neonicotinoid) that was first registered in

the United States in 1994 (NPTN 1998). It is used to control a wide variety of agricultural,

urban, public health, and veterinary pests and is estimated to account for 11-15% of the total

global insecticide market (Tomizawa and Casida 2005). Several formulations are available for

different treatment applications, only two are available for house fly control: Maxforce

Granular fly bait and Maxforce Fly Spot bait. Both products are baits, but differ from one

another by their formulation. Maxforce Granular fly bait is a red granule that can be applied as

a traditional scatter bait, within a bait station, or mixed with water and painted onto a surface.

The Maxforce Fly Spot bait, alternatively, is white wettable powder that is mixed with water

and sprayed onto a surface. When the Maxforce Granular fly bait is painted on a surface, or if

the granules become wet, it stains the surface red, whereas the Maxforce Fly Spot bait is clear

and does not stain.

Imidacloprid kills insects through contact and ingestion by agonizing the nicotinic

acetylcholine receptor (nAChR) (Fossen 2006). In house flies, imidacloprid is metabolized by

oxidation to the olefinn" metabolite, which has the same toxicological activity as imidacloprid

(Nishiwaki et al. 2004). Insects exposed to lethal doses of imidacloprid experience nervous

system excitability, modified feeding behavior, and death. Imidacloprid is considered a selective

insecticide because: (1) imidacloprid has a higher affinity for the insect nAChR's than

mammalian nAChR's, and (2) there are more nAChR's located in the insect nervous system than

what are found in mammalian systems (Yamamoto et al. 1995).









Resistance has yet to be reported in house flies (Gao et al. 2007), but it is well-documented

in Drosophila (Daborn et al. 2001). Cross resistance has been observed in multi-resistant house

flies (Wen and Scott 1997). Multiple resistance mechanisms are suspected in house flies.

Monoxygenase-mediated detoxication seems to be a primary mechanism in some strains of

house flies, but not in others (Wen and Scott 1997, Liu and Yue 2000).

Methomyl

Methomyl is a carbamate insecticide that was first registered in the United States in 1968

(EPA 1998). It is used to control a wide variety of agricultural, urban, public health, and

veterinary pests. Several methomyl formulations are available, but the 1% fly bait formulation is

the only one which is not classified as a restricted-use pesticide.

Methomyl causes mortality by contact and ingestion by inhibiting the acetylcholinesterase

(AChE) enzyme, which occurs in the central nervous system. Methomyl binds to AChE and

prevents it from binding to acetylcholine. This results in acetylcholine saturation at its neural

receptor, which results in a dramatic increase in generation of nerve impulses. Insects exposed

to methomyl show signs of hyperexcitability, convulsions, paralysis, and death.

Decarbamoylation of AChE is rapid and, therefore, carbamates are considered reversible AChE

inhibitors and recovery from sub-lethal poisonings can occur quickly (Yu 2007).

Little resistance to methomyl has been seen in house flies despite its frequent use and the

high levels of resistance seen in house flies to other carbamates (Barson 1989, Webb et al. 1989,

Scott et al. 2000, Darbro and Mullens 2004). Flies feeding on methomyl granules have been

found to receive a super-lethal dose that may play a large roll in why resistance has not been as

widespread (Price and Chapman 1987). Behavioral resistance, or bait aversion, has started to

become more apparent. In 1989, Barson (1989) reported that 8% of the resistant flies were

repelled by methomyl. In a study comparing the mortality of 35 field strains of house flies fed









methomyl in choice and no-choice tests, mortality decreased by nearly 30% when the flies were

given the choice test (Darbro and Mullens 2004).









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BIOGRAPHICAL SKETCH

Jeffrey Conrad Hertz was born in 1976 in Peoria, Illinois. His parents,

Marion Conrad "Butch" Hertz and Margaret Eloise "Weezie" Hertz (Hilton), raised him in

Lewistown, Illinois. They moved to Bernadotte, Illinois where he continued to attend school in

neighboring Lewistown until he graduated from Lewistown Community High School in 1994.

He entered the United States Navy and reported to basic training at Recruit Training Command,

Great Lakes, Illinois in November later that same year. Over the last 12 years, he served with

the United States Marine Corps, at Naval hospitals, and most recently, he was assigned to the

medical staff at the United States Capitol. He received his Associate of Science degree in

medical laboratory technology from George Washington University, Washington D.C. in 2002

and his Bachelor of Science in interdisciplinary studies, majoring in biology from Mountain

State University, Beckley, West Virginia in 2003. In 2004, he was selected as the very first

enlisted Sailor selected to study entomology under the Medical Service Corps In-service

Procurement Program (MSC-IPP). Upon graduation HM1 (FMF) Jeffrey Hertz will be

commissioned to the rank of Lieutenant Junior Grade as a medical entomologist in the Medical

Service Corps. He enjoys running and is an active member of Centennial Lodge #174 of Ancient

Free and Accepted Masons located in Upper Marlboro, Maryland. He, his wife, Karina, and two

children, Conrad and Kyra, are excited about their upcoming move to Jacksonville, Florida

where he will be working at the Navy Entomological Center of Excellence.





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1 POTENTIAL OF INSECTICIDE-TREATE D CORDS AND SPRAYABLE BAITS FOR CONTROL OF HOUSE FLIES (DIPTERA: MUSCIDAE) By JEFFREY CONRAD HERTZ A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2007

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2 2007 Jeffrey Conrad Hertz

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3 To my wonderful family

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4 ACKNOWLEDGMENTS I would like to thank the United States Navy for granting me the opportunity to pursue a Master of Science degree in entomology under the Medical Service Corps In-service Procurement Program: an experience I certainly didnt anticipate upon en listing. CAPT (ret.) Gary Breeden, CAPT Stan Cope, CAPT Mark Beavers, and LCDR Pete Obenauer provided exceptional support and irreplaceable mentorship th roughout this entire endeavor. All of which, I hope will continue throughout th e remainder of my career. Dr. Phil Koehler has been irreplaceable in his role as my academic advisor. His guidance and leadership through my entire term at the Univer sity of Florida has been exceptional. He has a knack for teaching that is unlike any other and his emphasis on extension and systematic problem solving will no doubt benefit me in my future role as a military entomologist. More has been gained from my committee, Dr s. Phil Koehler, Richard Patterson and Mike Scharf, in the past 2 years (espec ially the last 6 months) than ca n probably not be repaid in the next three decades. All have provided tremendous input not only on this current research, but on future work as well in both science and in lif e. All of which, is greatly appreciated and I sincerely hope that their i nput will not end upon graduation. Although I recently met Dr. Roberto Pereira, his infl uence will last the rest of my life. His drive for science and his skill at teaching is inspiring and his s upport has been nothing less than remarkable. He would always find time to lis ten, critique, review, and analyze anything I brought to him despite his own hectic schedule and workload. My research would not have even begun w ithout the tremendous work by the Urban Entomology Laboratory Fly Group and the generosity of Dr. Phil Kaufman for allowing the fly group to use his laboratory space. Ricky Vazquez or ientated me and gave me the tools needed to continue his successful fly program. Dr. Matt Aubuchon, Ryan Welch, Terry Krueger, and Mark

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5 Mitola made the fly program function every day ove r the last two years a nd without all of their consistent work the colony would have surely cr ashed. Terry Krueger provided more help than any other when it came to my research. He wa s always there to provide a helping hand and would often do recordings when I was unavailable to do so. Without his help, this research could not have been completed. Two fraternal brothers helped in every way they could th roughout my University of Florida experience. Tiny Willis kept me complete ly stocked and ready to go at all times and was always a pleasure to talk to a bout life, politics, and the good ol days whenever I needed a break from my research, writing, or class. HM1 (SCW ) Joseph W. Diclaro II, is an old friend who seems to follow me wherever I go and I appr eciate that because it seems good things come whenever he does. Hopefully his time at the Univ ersity of Florida will be as memorable as mine. So mote it be. Many new friends have been made and their fr iendship is valued. To mention them all would cumbersome. The entire urban entomology lab and Urban Entomological Society have all been great to work with a truly extraordin ary group of scientists. In addition, the whole administrative staff in the Entomology Department has been outstanding. If it wasnt for them I can say with complete confidence that my gradua tion, as well as many others, would not come to pass. Finally, my family has been the foundation for th is entire chapter in our lives together; my wife, Karina, the cornerstone. Never once did sh e question time allotted away from her or our two children, Conrad and Kyra, for or was sh e ever not willing to edit manuscripts when obviously the subject matter did not suit her interests. Conrad and Kyra deserve a very, very special thank you for being such really great children always Daddys little helpers.

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6 TABLE OF CONTENTS page ACKNOWLEDGMENTS...............................................................................................................4 LIST OF TABLES................................................................................................................. ..........8 LIST OF FIGURES................................................................................................................ .........9 ABSTRACT....................................................................................................................... ............10 CHAPTER 1 STATEMENT OF PURPOSE................................................................................................12 2 REVIEW OF LITERATURE.................................................................................................14 Classification, Origi n, and Distribution..................................................................................14 Identification................................................................................................................. ..........14 Egg............................................................................................................................ .......14 Larva (maggot)................................................................................................................14 Pupa........................................................................................................................... ......15 Adult.......................................................................................................................... ......15 Sex Differentiation..........................................................................................................16 Life Cycle..................................................................................................................... ..........16 Nutrition, Longevity, and Overwintering...............................................................................17 Flight, Movement, and Resting Behavior...............................................................................19 Pest Status and Health Importance.........................................................................................21 Control........................................................................................................................ ............23 3 INSECTICIDE-IMPREGNATED CORDS FOR HOUSE FLY CONTROL........................29 Introduction................................................................................................................... ..........29 Materials and Methods.......................................................................................................... .30 Insects........................................................................................................................ ......30 Laboratory Arenas...........................................................................................................31 Cord Attractiveness Bioassay..........................................................................................31 Impregnated-Cord Laboratory Bioassays........................................................................32 Impregnated-Cord Field Cage Bioassay..........................................................................33 Statistical Analysis..........................................................................................................34 Results........................................................................................................................ .............35 Discussion..................................................................................................................... ..........37 4 EVALUATION OF A NEW IMIDACLOPRID BAIT FOR HOUSE FLY CONTROL......46 Introduction................................................................................................................... ..........46 Materials and Methods.......................................................................................................... .48

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7 Insects........................................................................................................................ ......48 Laboratory Arena Design................................................................................................48 Field Cage Design...........................................................................................................49 Fly Bait Comparisons......................................................................................................49 Bait-Treated Cords..........................................................................................................51 Data Analysis.................................................................................................................. .52 Results........................................................................................................................ .............52 Fly Bait Comparisons......................................................................................................52 Bait-Treated Cords..........................................................................................................53 Discussion..................................................................................................................... ..........54 5 SUMMARY AND CONCLUSIONS.....................................................................................64 APPENDIX 1 REVIEW OF INSECTICID E-IMPREGNATED CORDS.....................................................67 2 REVIEW OF FLY BAITS......................................................................................................70 3 REVIEW OF INSECTICIDES EVALUATED......................................................................72 Fipronil....................................................................................................................... ............72 Indoxacarb..................................................................................................................... .........73 Imidacloprid................................................................................................................... .........74 Methomyl....................................................................................................................... .........75 LIST OF REFERENCES............................................................................................................. ..77 BIOGRAPHICAL SKETCH.........................................................................................................85

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8 LIST OF TABLES Table page 3-1. Efficacy of various cords impregnated with 0.1% fipronil or 0.6% indoxacarb on female house flies............................................................................................................. .40 3-2. Cumulative number of dead flies and percent fly count reduction in relation to control fly counts of house flies expose d to 0.1% fiproniland 1.2% indoxacarbimpregnated cords in field cages........................................................................................41 4-1. Number of dead and percent fly count re duction in relation to control fly counts of house flies exposed to imidacloprid bait-tr eated lattice squares in field cages.................59 4-2. Number of dead and percent fly count re duction in relation to control fly counts of house flies exposed to imidacloprid ba it-treated cords in field cages...............................60

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9 LIST OF FIGURES Figure page 3-1. Laboratory and field experi mental design elements..........................................................42 3-2. Attraction of female house flies to various natural and synthetic cords............................43 3-3. Female house fly mortality exposed to various natural and synthetic cords treated with 0.1% fipronil for 24 h (A) a nd 0.6% indoxacarb for 48 h (B)...................................44 3-4. Comparison of the most attractive cord (m anila) and the least attractive cord (nylon parachute) in the cord at tractiveness experiments.............................................................45 4-1. Mortality of female house flies exposed to imidacloprid and methomyl granular scatter baits and a sprayable imidacloprid bait..................................................................61 4-2. Morbidity (knockdown) of female house flie s exposed to natural and synthetic cords dipped in a 2.5% solution of imidacloprid sprayable bait.................................................62 4-3. Mortality of female house flies exposed to natural and synthetic cords dipped in a 2.5% solution of imidacloprid sprayable bait....................................................................63

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10 Abstract of Thesis Presen ted to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science POTENTIAL OF INSECTICIDE-TREATE D CORDS AND SPRAYABLE BAITS FOR CONTROL OF HOUSE FLIES (DIPTERA: MUSCIDAE) By Jeffrey Conrad Hertz August 2007 Chair: P.G. Koehler Major: Entomology and Nematology House flies are often controlled using insecticides when the s ource of infestation can not be located or remedied by non-chemical methods Historically, hous e flies have shown a tremendous potential to develop insecticide resistance and with few classes of insecticides currently registered for house fl y control, new products and methods need to be evaluated to prevent future control failures. This resear ch evaluated the potent ial use of two innovative methods to control house flies: fipronila nd indoxacarb-impregnated cords and a sprayable imidacloprid fly bait. For the insecticide-impregnated cord studies eight various natural and synthetic cords were evaluated to determine which cords were attractive to house flies. Natural cords were more attractive than synthetic cords; the plant-based manila cord was most attractive and the nylon parachute cord was least attractive. The most attractive cords (manila cotton, wool, nylon, and polypropylene) were treated with 0.1% fiproni l or 0.6% indoxacarb and evaluated in the laboratory to determine their effectiveness. A ll cords were more effective than the impregnated cotton cord except the fipronilimpregnated nylon cord (LT90) and the indoxacarb-impregnated polypropylene cord. The wool cord was the most effective, LT50 (Fipronil = 12.9 h; Indoxacarb = 32.6 h) and LT90 (Fipronil = 22.4 h; Indoxacarb = 51.5 h) The wool cords were impregnated

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11 with 0.1% fipronil and 1.2% indoxacarb and evalua ted in a controlled field environment with fresh cords and cords that were aged 4 wk. No significant differences were seen between fly count reductions of either treatment. Both treatments reduced fly c ounts by >57% by 24 h and >87% by 48 h with both fresh and aged cords. A reduction in efficacy was seen with aged cords. The new imidacloprid sprayable fly bait form ulation was compared against two commonly used dry scatter baits in the laboratory and ag ainst a granular imidacl oprid paint-on bait in a controlled field setting. Additiona lly, the sprayable bait was evaluated for use in impregnated cords. No differences were seen in mortality be tween the three scatter baits in the laboratory or between the imidacloprid baits in the field cages. Both imidacl oprid baits reduced fly counts by >70% in the field within 24 h, but were not effective after treatmen ts were aged for 2 wk. When various cords were treated with 2.5% of the new bait, the wool cord had higher mortality (74%) compared to the other natural and synthetic cords tested. Knockdown recovery was observed with all bait-treated cords in the laboratory, but was not determined to occur in the field cages. The bait-treated cords reduced fl y counts by >82% with fresh cords and cords aged for 4 wk. Impregnated cords and the new sprayable ba it should prove to be valuable tools in established fly management programs in urban, ag riculture, and military settings. Fipronil and indoxacarb are not currently registered for hous e flies, but both appear to be effective insecticides for their control.

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12 CHAPTER 1 STATEMENT OF PURPOSE Throughout history flies have undoubtedly been a nuisance to both man and animal alike; however, because of their propensity to frequent pathogen-rich filth they do pose a human health risk. House flies have been shown to tran smit numerous pathogens and their synanthropic behavior may make house flies one of the most troublesome insect vectors (West 1951, Greenberg 1973). This is especial ly true in areas affected by natu ral disasters or conflict. Often times following these chaotic events, basic sanita tion measures are out prioritized for casualty recovery and, as a result, tremendous populations of house flies emerge. Chemical insecticides are often used in thes e situations or any s ituation where rapid house fly control is needed. Today there are more chemi cals registered as insectic ides than ever before, but few of these insecticides are registered for house fly control. The insecticides that are registered for house fly control only come fr om five chemical classes: organophosphates, carbamates, pyrethrins/-oids, triazines, and neonicotinoids. The organophosphates and the carbamate insecticides continually get furthe r restrictions by the Environmental Protection Agency (EPA) limiting their use and their future availability in th e United States may be bleak. In addition, house flies have consis tently shown the ability to deve lop resistance to all chemicals used to kill them (Liu and Yue 2000, Scott et al. 2000, Kaufman et al. 2001) and having only 3-5 chemical classes to rotate with may prove to be detrimental to a fly management program. There is an immediate need for new insecticides regist ered and new techniques for house fly control to prevent future control failures. In 2004, the Department of Defense (DOD) established the Deployed War-Fighter Protection (DWFP) program to deve lop and test management tools for pest and vector species, including house flies, which tran smit diseases to the deployed wa r-fighters. The Armed Forces

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13 Pest Management Board (AFPMB) administers the DWFP program and specifically requested research to improve or devel op integrated filth fly control strategies and non-conventional pesticide methodologies. Insecticide-impregnated cords and sprayable fly bait may be beneficial tools that the deployed war fight er could use for fly manageme nt programs. The research contained herein was designed to provide new information regarding these techniques to the DOD.

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14 CHAPTER 2 REVIEW OF LITERATURE Classification, Origin, and Distribution The house fly, Musca domestica belongs in the class Hexapo da, order Diptera, suborder Brachycera, infraorder Muscomorpha (Cyclorr hapha), and family Muscidae (Triplehorn and Johnson 2005). It was first described by Linnaeus (1758). It is believed the family Muscidae evolved sometime during the Permian period of the Paleozoic era (Lambrecht 1980). The exact origin may never be known, but many speculate that the house fly originated in the Middle East ar ea of the Palearctic re gion (Skidmore 1985, Pont 1991) and was distributed through multiple intr oductions into the New World (Marquez and Krafsur 2002). Today, house flies are one of the mo st commonly found synanthr opic pests. It is found in virtually every region of the globe that man or animal exist. The only exception is areas, such as high altitudes and the arctics, whic h are prone to extreme cold temperatures (West 1951). Identification Egg House fly eggs are white, bluntly rounded, banana-shaped eggs approximately 1 mm in length, and often laid in clusters (Keiding 1976). The egg widens in size posteriorly to anteriorly and the dorsal surface has two longitudinal, curved ridges that narrow just prior to reaching the caudal end. Larva (maggot) House flies have three larval instars. E ach instar has no eyes legs, antennae, or appendages and are commonly known as maggot s (Moon 2002). The maggot has a rounded posterior that tapers to a point towards its head. A pair of bl ack spiracular plates is located

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15 posteriorly, which progressively becomes more chitinized and D-shaped through molts. First and second stage larvae have two spiracular openings (slits) used for gas exchange and a third opening appears on the third instar larvae (Moon 2002). Prothoracic spiracles are fan-shaped and appear after the first molt. A cephalophary ngeal skeleton, comprised mainly of sclerotized mouth hooks, is located at the anterior end of the larvae. Pupa House fly pupae are approximately 6.3 mm in length (West 1951). At the beginning of pupation, the puparium is white in color but eventually become s reddish-brown. The puparium is medially enlarged with bluntly rounded ends. Two pupal horns are locate d laterally just prior to the posterior boundary of the first abdominal segment (Siriwattanar ungsee et al. 2005). Posterior spiracles are located on the posterior end and appear as two flat, circular prominences. The anterior spiracle is situated on the puparium in the same locati on as in the thir d instar larvae (Siriwattanarungsee et al. 2005). Adult The adult house fly is a medium-sized (6-9 mm) gray insect with large brown compound eyes (Moon 2002). On the vertex, between the eyes, lies the ocellar triang le containing the three simple eyes. The house flys antennae are also located between the eyes within the triangular facial depression. The antenna is six segmented, but only appears to be four. The first three segments, the scape, pedicel, and large first flage llar segment, give rise to the three-segmented arista. Segment one and two of the arista is ambiguous; segment three is bristlelike. The sponging proboscis of the house fly terminates to a heart-shaped sucker. The proboscis can be greater than the length of the head when fully extended or obscure when fully retracted. Two brownish-black maxillary palpi lie on the anterior margin of the proboscis.

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16 The thorax of the house fly has four black long itudinal stripes that ca n be viewed dorsally. Attached laterally to the mesothorax are tw o membranous wings. The wings, when extended, are approximately twice the distance of the fly s length. At rest, the house fly pulls the wings back incompletely over the abdomen forming an overall triangular appearance from above. The fourth longitudinal wing vein shar ply angles towards the wing ape x. Situated below each wing is a knob-shaped organ used for equilibrium called th e haltere. The legs of the house fly attach ventrally to each segment of the th orax and all legs have five-segmented tarsi. The first tarsal segment is much longer than all other segments and the fifth segment bears two claws, a hair-like empodium, and a sticky pad called a pulvillus. The abdomen is gray, dorsally, and cream-color ed ventrally. Five pa irs of spiracles line the ventral surface of the female; six pairs line the ventral surface of the male. The tip of the abdomen ends in either the scle rotized genitalia of the male or the retracted ovipositor of the female. Sex Differentiation Adult female house flies are almost always la rger than adult males. Additionally, males can be differentiated from adult females by locati ng the dark sclerotized genitalia plate located on the distal aspect of the abdomen. The tiny ma rk made by the ovipositor tip of the female is very distinctive compared to the male geni talia especially when females are gravid. Furthermore, adult house flies can be separate d by the gap distance that divides the compound eyes. Females have a much wider space separating the eyes when compared to male counterparts. No differentiation can be made in the immature stages. Life Cycle Male and female house flies can successfu lly mate 24 hours after emergence from the pupae (Murvosh et al. 1964). Prior to copulatio n, a male will seize a resting female or strike a

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17 flying female at which point they fall to a surface. If a copulating pair is disturbed while mating they may attempt to fly a short distance to an a lternate surface. Copulat ion can last for more than 1 h, but sufficient sperm transfer can occur in less than 10 min (Murvosh et al. 1964). Once successful copulation takes place, the female is fertilized for life. Batches of up to 150 eggs are laid 4-8 days after copulation (West 1951). The fe male house fly carefully embeds her eggs into practically any fermenting organic material The eggs hatch within 24 hours, and 1st instar larvae emerge and begin to feed (West 1951). The la rvae undergo two molts within 3-5 days before pupation (Hogsette 1995). P upation begins when the 3rd instar larva stops feeding and constricts within its own integument. This makes a whit e puparium which turns reddish-brown within 24 hours. After 3-5 days, the adult breaks thr ough the anterior end of the puparium using a temporary, inflated sac located on its head calle d the ptilinum. Once free from the puparium, the newly emerged adult house fly hops around to le t its wings extend and cuticle harden. Nutrition, Longevity, and Overwintering House flies larvae have been reared in the la boratory on practically every type of filth imaginable. Today, they are most often reared in a medium containing animal feed and water (Hogsette 1992). Fermenting odors attract gravid females to oviposit on breeding sites in the field, but understanding precise ly what nourishes a maggot with in the medium is not fully understood. All house fly maggots are saprophagous a nd feed on liquids or substrates that are readily dissolved by droplet regur gitation (Nation 2002). It was originally thought that bacteria were essential in the development of house fly maggots, however many have successfully reared them in aseptic media (Brookes 1956, Monroe 19 62). Despite this, bact eria still may have provide nutritional value (e.g. vitami ns) to maggots (Zurek et al. 2000). Adult house flies are omnivorious and emerge with little stored energy and nutrients (Moon 2002). They begin to feed within 2-24 hou rs after emergence (Keiding 1976). In order to

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18 survive, they must find a sugar source, or othe r assimilable starch, and water (West 1951). In addition, female house flies require a pr otein source for vitellogenesis. When feeding, adult house flies are attracted fi rst visually, then when they are within a detectable range, by smell using their antennae (Keidi ng 1976). Flies locat e the source of the aroma by smelling the substrate with chemoreceptors located on the lateral aspect of their 2-5 tarsi. Once their tarsi are in contact with a suit able substance, the fly extends its proboscis and begins to feed. Liquid substances can be r eadily imbibed, but solids are ground down using the prestomal teeth on the proboscis and emulsified using a vomit drop originating in the crop and salivary glands. The largest part icle a house fly can ingest is 40 (Greenberg 1973). Ingested liquid and emulsified particle s enter the pseudotracheae and th en pharynx. Once past the pharynx, liquids pass into the crop and emulsified food particles enter the proventriculus then the ventriculus. The crop is connect ed to the pharynx by a long sle nder tube lined with numerous sphincters that controls the flow of liquid back to the abdomen wh ere the bilobed crop is housed. The heamolymph osmotic level dictates the rate th e crop empties into the ventriculus. The more concentrated the sugar meal, the slower the cr op empties (Greenberg 1973). The ventriculus empties into the longest part of the alimentary track, the proximal intestine. The proximal intestine is divided from the di stal intestine by excretory organs called the Malpighian tubules. The distal intestine terminates at the anus. Longevity of any organism can be highly va riable. Food availability, environmental conditions, and activity of an individual fly greatly influences how long it will live. Of the three survival-mandated nutrients, sugar is the most critical for survival. Lysyk found that the availability of sucrose was the most important factor promoting longevity, followed by other food sources (manure, milk) and temperature (1991). Flies will live 50% longer on sucrose

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19 alone, than they do on water alone (Greenberg 1960). However, f lies without water generally die within 48 hours (West 1951). Temperature is inversely proportional to th e life span of the a dult house fly; higher temperatures reduce the life expectancy, while lo wer temperatures increase it (West 1951). In laboratory conditions where adequate food is provided ad libitum and environmental conditions are controlled, male house flies can live up to 40 days and fema le house flies can live up to 60 days (Rockstein 1957). In the field, house f lies are estimated to only live about 10 days (Hogsette 1995). Bucan and Sohal (1981) found that adult males and females isolated from the opposite sex live longer than when they are housed together. To increase their survivability when temper atures drop below optimum levels, house flies survive by overwintering in buildings and animal c onfinements. All life stages are susceptible to subzero temperatures, so microclimates must exist that allow flies to propagate. Rosales et al. (1994) concluded that house flies require habitats that are above -5C, and must stay above 10C long enough for the house fly to complete its life cycle. Flight, Movement, and Resting Behavior Flies have two wings located on the lateral aspe ct of the body on the pteropleura. Directly above the metathoracic coxae are the vestigial wings, or halteres, which are used as gyroscopes for equilibrium. Like other flying insects, a house fly achieves flight by creating wing movement through indirect thorax compression and decompression cau sed by the flight muscles. These flight muscles comprise approximately 11% of the total body weight in the genus Musca (Greenberg 1973). This musculat ure makes house flies extremely st rong fliers and very capable of flying upwind in mild and moderate winds. Fly movement can be classified as dispersal, dispersion, or migration (Greenberg 1973). Dispersal is any active movement within a relati vely small defined area. House fly problems are

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20 often localized near a source of infestation (Howard 2001, N azni et al. 2005). Often times, dispersal will be dependant on the sun light. Flies tend to follow the sun and more will be located where the sun is shining (Anderson 1964). Th is is especially true in cooler temperatures; in hot temperatures, flies may avoid the sun and s earch for cooler locations. Dispersion is the movement of flies between adjace nt areas and often involves the movement assisted by passive transport. Passive transport can occur on garbag e trucks, tractors, auto mobiles, or any other vehicle including strong winds. This is often se en when breeding sources are sporadic or when no breeding sources are near and flies move into th e area in search of new oviposition sites. This movement is why flies are found in areas where no apparent fly breeding material is present. Migration is any directed and su stained flight that often occurs seasonally. This type of movement is not normally associated with house f lies, however, many have been trapped in areas that would suggest that migration was the on ly possible explanation (West 1951, Jones et al. 1999). Passive transport may also play a large role in these situations. Fly movement can be influenced by many fact ors such as odors, wind, weather, time of day, and population structure. F ood and oviposition sites are probabl y the most critical factors (Bishopp and Laake 1921). However, many questions still need to be answered on what is attractive or needed by house flies since flies will often leave one oviposition site in search for an alternative site although sources are immediately available an d sufficient for survival. When seeking these new food sources or oviposition sites, flies can fl y upwind in mild to moderate wind speeds, but strong winds, or even shifting winds, can disperse house flies to areas where survival may not be suitable. Taylor (1974) found that house flies are day f liers and their flight activity increases with high temperatures (25-30 C) and low humidity (50-65%). Flies always

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21 seek temperatures above 15.5C, but are capable of flight at temp eratures below 55C (Greenberg 1973). House flies have distinct resting behavior. In warmer climates, flies prefer to rest at night outdoors on low hanging twigs of trees and bushes, but may still be seen resting indoors on wires or cords close to the ceiling (S cudder 1949, Keiding 1965). In cold er climates, house flies will rest exclusively indoo rs. In all cases, flies tend to rest on ob jects with distinct edges less than 4.5 m from the ground, shielded from direct wind (Scudder 1949). Keiding and Hannine (1964) found a distinct preference for hous e flies to rest on objects suspe nded vertically from ceilings, however, Fay and Lindquist (1954 ) found no differences in orientation of suspended cords. The visual orientation of house flies to objects has been widely disput ed. Objects that are light in color, smooth, or metall ic are highly avoided by flies; wh ereas, objects that are dark in color and rough are generally more frequently rested upon by flies (Areva d 1964). Hecht et al. (1968) performed a number of indoor and outdoor experiments to determine the attraction of house flies to different colored cardboards. They found that black was most preferred indoor and white was the most preferred outdoor. When combining the indoor and outdoor results, the red colored surfaces were most preferred. The leas t preferred colors were blue (indoor) and brown (outdoor). The attraction to the white surface out doors was attributed to a flys attraction to ultraviolet light because of the reflective qualiti es of the white cardboard. Flies see wavelengths between 350-480 m (McCann and Arnett 1972). Contrast s of colors (dark on light/light on dark) may be very important to the attraction of house flies. Howard and Wall (1998) counted more flies on white surfaces with black backgrounds than on any other black/white combination. Pest Status and He alth Importance House flies are renowned for thei r ability to annoy anyone and anything they are near. It only takes one house fly to turn a customer away from a restaurant and only a few to disrupt

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22 production and morale at a work site. Most s ee house flies as a sign of unhygienic conditions and attempt to avoid them at all costs. Flies leave fecal and vomit spots on work equipment, consumables, and personal items most of whic h are not be generally quantified in economic losses but their impact can easily be seen when comparing two similar establishments one with a fly problem and the other lacking a problem. L itigation cases due to house flies have increased in the United States recently due to the migra tion of urban dwellers d eeper into rural settings where livestock and poultry farms have a relative ly large abundance of flies a situation many urban dwellers may not be familiar with. Although primarily nuisance pests, house flies do pose a risk to the health and well-being of man and livestock. Because of a house flys beha vior and survival needs, it frequently comes in contact with pathogenic organisms. Hous e flies are extremely capable of transmitting pathogens mechanically (West 1951). West (1951) and most recently Greenberg (1973), have compiled extensive lists of the pa thogens (bacteria, viruses, fungi protozoa, and nematodes) the house fly is capable of tran smitting. The transmission of Campylobacter spp. E coli O157:H7, H. pylori, C. parvum, and G. lamblia are probably the most significant pathogens capable of being transmitted by the house fly recently repo rted (Shane et al. 1985, Grubel et al. 1997, Kobayashi et al. 1999, Graczyk et al. 2003). Of particular importance is the exponential pro liferation of house flies following situations arising from natural disasters or conflict. Following the tsuna mi that devastated parts of Indonesia in 2004, sewage and draina ge systems were destroyed l eaving sewage pools that bred numerous species of filth flies (Burrus 2005). This is often what occurs following any of these chaotic events; communities are left in shambl es and multiple oviposition sites develop due infrastructural collapse or basic municipal sanita tion being out prioritized. Large fly populations

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23 then develop and epidemic levels of dia rrheal cases normally follow (Thornton et al. 2005, Watson et al. 2007). This reduces military readine ss and stresses health care systems (Putnam et al. 2006). The direct impact of house flies on dis ease transmission in these situations is either not measured or often not measurable because the same pathogens transferred by house flies can be just as easily transferre d by man or other organisms. Control The house fly is best controlled through a fly management program based on the sound principles of Integrated Pest Management (IPM), including a comb ination of monitoring, cultural, biological, and chemical control meas ures (West 1951, Keiding 1976). Monitoring is any technique employed to determine presence/ absence and peak/trough flows of house fly populations. Cultural controls are any measures that deliberately alter the life cy cle of the house fly without the use of chemical or biological agents Biological controls are agents or organisms that alter the life cycle of the house fly and chemical contro ls are naturallyor syntheticallyderived chemicals that can alter the life cycle of the house fly. Every aspect of the fly management program s hould have some form of fly monitoring to determine when and what type of approach shoul d be employed (pre-treatment survey) and to see the effectiveness of the treatment (posttreatment survey) (Keiding 1976). In the pretreatment survey, house fly densit y, distribution, and beha vior should be noted to help determine which treatment option to use. Post-treatment surveys normally only need to monitor fly densities unless failure occurred. In this ca se, a complete reassessment should be done to include some form of monitoring for insecticide resistance. There are four basic methods to obtain a fly population index: counti ng flies, counting fly specks, netting flies, or trappi ng flies (Keiding 1976). If counti ng adult flies, several methods can be used. The classic technique is the use of a Scudder grill a simple grid constructed of

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24 wood that can be placed over an infestation sour ce and then all flies landing on the grid are counted (1947). A similar technique that can be used in practically any situation is to simply mark an area (preferably near infestation or re sting areas) and count flie s landing on it. Counting fly specks left on index cards may be the method of choice today for indoor sampling because of its simplicity. Spot cards can be positioned in standard locations throughout the infested area and will give a good representation of the fly popul ations over time. Simply hang them and check back on them after a designated time peri od. Over time it will show peak and troughs of fly populations; in addition, the spot cards can be archived and marked with any insecticide treatment used to provide a dditional documentation for resistance monitoring. Some users, however, prefer to use destructiv e sampling. In these cases, baited traps, sticky ribbons, or even netting flies can work well. All of the above methods work and can give consistent numbers indoors as long as the same sampling method is used for all counts. Ho wever, infestations that occur outdoors are not as easily monitored. With outdoor sampling, s pot cards are an unrealistic method because placement would be difficult and precipitation woul d destroy them. Scudder grills and other fly count techniques are subject to wide variation depending on positio nal effects, time of day, and weather conditions (Geden 2005). Baited traps and net sweeping ma y be the best techniques to use outdoors for surveillance work, but these methods are destructive and not suitable in every situation. Beck and Turner (1985) found that usi ng a simple visual index correlated better with absolute fly densities than spot cards, sticky ribbons, scudder grill and fly counts, but these indexes are very subjective and will vary between persons making the counts. Perhaps, the best way to monitor house flies outdoors is to use a combination of the methods described.

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25 By far the most important aspect of a fly management program is the use of cultural controls. Cultural controls target the breeding and feeding sites of adul t and larval flies. Additionally, they are used to prevent adult flies from c ontacting food, pathogens, and man (Keiding 1976). Garbage is the main source of infestation in an urban environment and manure is the main source in an agri culture environment, however, bot h sources can be found in any environment. In developed urban communities, th ese breeding sites are normally controlled by very established municipal san itation measures (e.g., closed se wers, garbage removal, etc.) (Hogsette 1995). Agriculture facilities have to physically remove manur e or bake it by covering dung heaps with plastic sheeting. Garbage should be removed from the area at least twice weekly or burned (Keiding 1976). Windows, screen s, and doors should all be in good repair and kept closed to prevent flies from entering esta blishments. The installa tion of air curtains on doors and windows that are frequent ly opened and closed is an en ergy efficient option that can reduce the number of f lies that enter. Traditionally baited traps, light traps, electrocuting light tr aps, and sticky ribbons have been used as cultural control measures, but th eir effectiveness at re ducing house fly populations are limited and their use should primarily be c onsidered as a monitoring technique. However, these methods do trap and kill flies so using them should never be automatically ruled out. In fact, if the likelihood of large infestations does no t exist, then traps are a good method for killing flies (such as in grocery stores and restaurants) but some consider ations should be made prior to their use. Baited traps generally can not be us ed indoors or near residences because the odor associated with this method is repulsive to hum ans (Pickens et al. 1973) Electrocuting light traps release fly parts, bacteria and viruses and may be just as unhygienic as the fly itself and should not be used in areas were conditions need to remain relatively aseptic (Urban and Broce

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26 2000). Lighted traps need bulb replacement appr oximately every six months, and sticky ribbons need replaced frequently due to dust and fly cadaver build -up. Flies that do make their way into an establishment can be physically remove d by numerous devices, but the most common physical cultural control method is the good ol fash ioned flyswatter. A novelty gadget used for killing insects, including flies, has recently sparked numerous videos on the World Wide Web. The device is a combination electrocuting trap and fly swatter. This device may be interesting and fun, but the same risk is associated with it as the regular el ectrocuting light traps. The second most important principle of a good fly management program is the use of biological controls. Every stage of a flys life cycle is vulnerable to attack by some form of biological control (West 1951). The eggs are of ten predated upon by mites, earwigs, ants, and some beetles. Larvae are also attacked by mites, earwigs, and beetles, as well as some birds, wasps, and other Dipteran larvae. Pupae are of ten parasitized by small wasps and some beetles, others can be eaten by birds and large beetles. Many adult house flies meet their demise thanks to predatory insects (mantids, flies, dragonflies, wasps, ants) and arachnids. Many other adults are eaten by reptiles, amphibians, small mammals, and birds. House flies are also prone to infections by bacteria and fungi. Fortunatel y, all of these biological controls are already abundant in a flys natural envi ronment and the goal of a fly management program should be to maintain or supplement these existing populations (Geden 1995). Parasitic wasps can now be purchased commercially and their successf ul use is variable (Axtel 1999). The use of chemical insecticides is th e third component of a good fly management program. Chemical insecticides provide quick resu lts (i.e. dead flies) an d satisfactory control in one or two days, but their use should be limited to reduce the likelihood of resistance evolution and maintenance of non-target organisms. Chemical insecticides can be applied several different

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27 ways: residual surface treatments, larviciding, space sprays (including aerial spraying), and baits. The most common method for fly control is the use of space sprays and dry insecticidal baits. Residual surface treatments can be applied to any surface in any location the label allows but they are most effective when applied direc tly to fly resting areas (Keiding 1976). Neglecting to monitor and treat areas where flies are primarily resting will result in excess insecticide usage and population reductions of non-target organisms. Several insecticid es are available for residual treatments; most are organophosphate based. All are generally good for long term control, but there excessive use may increase sele ction for insecticide resi stance. One technique for residual insecticide application is the use of insecticide-impregnated cords which target the distinctive behavior of flies to rest on objects with edges (S cudder 1949, Keiding 1965). This method is thought to be less likely to select for resistance because the treated area is small and treatments can be readily removed or replaced w ith additional cords treated with insecticides from different chemical classes (Appendix A). Larviciding with insecticides sounds great in theory because larvae are relatively nonmobile compared to the adult flies, which have th e ability to fly to differe nt areas if one is found unsuitable; however, larviciding is really not a practical method fo r extended control. Larvicides have to be applied frequently because they ar e applied to areas such as garbage and manure both of which constantly accumulate. Larvicides kill non-target organisms coming in to feed on the house fly immature stages and would reduce t hose populations over time. In addition, if the same class of insecticide is used for larviciding th at is used for adult cont rol resistance selection would be rapid. Larvicides are e ffective if sites to be treated are expected to exist for a short period of time; in these instan ces several organophosphate ins ecticides and insect growth regulators can be used.

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28 Space sprays are primarily pyrethrinor pyrethr oid-based insecticides, but some may also be organophosphate-based, that ar e sprayed into the air in a fl y infested space or over a fly infested area. These types of insecticides target the fly nervous system and cause rapid knockdown of contacted flies (Yu 2007). Space spra ys are more effective when an abundance of flies are concentrated in one area; this occurs mainly in th e evening indoors and in the morning outdoors (Keiding 1976). They have little to no re sidual and have to be reapplied frequently (daily) in areas of large infestat ions. Another type of space treat ment is made through the use of insecticide vaporizers. The only vaporizer currently available is formulated with the organophosphate dichlorvos, but its use is becoming more restricted and its future longevity may be short lived. Insecticide baits are easy-to-use insecticides that have added attractants into the formulation matrix to draw flies into the treated area to contact the insecticide either by ingestion or contact. A basic bait matrix is a simple solution of sugar, water, and an insecticide. Complex bait matrices contain multiple sugars, pheromones (Z-9-tricosene), and other substances found attractive to house flies. The most widely used fl y baits available are form ulated in dry granules as scatter baits containing carbamates and ne onicotinoids (Appendix B), however, other bait products are available and frequen tly used. Like space sprays and larvicides, baits have to be frequently applied because environmental conditi ons degrade them or they become covered by manure or garbage. Also, when baits are used in areas with large fly populations, the flies will consume the bait rapidly leaving li ttle bait behind for the immature stages that will eventually emerge.

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29 CHAPTER 3 INSECTICIDE-IMPREGNATED CORDS FOR HOUSE FLY CONTROL Introduction The house fly, Musca domestica L., is widely considered the most common nuisance pest. Their nuisance pest status can quickly change to a public health risk if fly populations occur near inhabited areas where pathogen-ri ch oviposition sites are found. Areas stressed due to natural disasters, humanitarian crises, or combat are of ten plagued by large fly populations (Rosales and Prendergast 2000, Burrus 2005, Thornton et al. 2005). The most effective way to control house flies and reduce the risk of disease transmission is by eliminating their pathogen-rich oviposition sites. Oviposition site removal may be imprac tical, especially in areas affected by natural disasters and combat, where th e oviposition sites are too numer ous or difficult to reach. The best control method to use when sanitati on fails or when fly populations need to be rapidly controlled are chemical insecticides. Chemical insectic ides provide rapid kill of house flies and markedly reduced fly densities can be achi eved in as little as 12 days. Baits and space sprays are the primary chemical insecticide me thods used for house fly control today, but both methods provide little or no resi dual control, and resistance to their active ingredients is well documented in house flies (Georghiou and Lagune s-Tejeda 1991, Liu and Yue 2000, Scott et al. 2000). In addition, the number of registered inse cticides available for house fly control in the United States continues to decrease (Kaufman et al. 2001). New insecticides and application methods are clearly needed to avoid futu re insecticide resistance problems. Insecticide-impregnated cords have been used with great success to control flies and are considered less likely to select for resistance than traditional residual sprays (Keiding 1976). Their first use was in 1947 (Baker et al.) a nd by the mid-1950s, insecticide-impregnated cords were commercially available and widely us ed (Fehn 1958, Smith 1958). The commercially

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30 available cords contained 13.79% parathi on and 3.54% diazinon (Smith 1958). Cords impregnated with high concentrations (up to 25% active ingredient) of other organophosphate and organochlorine insecticides were also widely used with great success (Kilpatrick and Schoof 1959, Keiding 1976, Rabari and Patel 1976). These products are no longer used today due to the popularity of insecticidal baits and space spra ys and because the Environmental Protection Agency, acting under federal legislation, eliminat ed the use of their active ingredients. The objective of this study was to investigate if cords impregnated with newer insecticides would be an effective tool for house fly contro l. Specifically, the objectives were to: 1) determine the attractiveness of va rious natural and synthetic cord s to house flies, 2) determine the effectiveness of fipronil and indoxacarb on the mo st attractive cord materials, and 3) evaluate the effectiveness of the best cord/treatment combination in a simulated field environment. Materials and Methods Insects. The Horse Teaching Unit (HTU ) strain of house flies, M. domestica L., reared at the University of Florida in Gainesville was used for all experiments. Larvae were reared on a diet medium, modified from Hogsette (1992), containing 3 liters wheat bran, 15 ml methyl paraben, 1.5 liters water, and approximately 200 g (250 ml) dairy calf feed (Calf Manna pellets, Manna Pro Corp., St. Louis, MO). A ll developmental stages were held at 26 1 C and 55% RH with a 12:12 (L:D) photoperi od. Adult flies emerged within screened rearing cages and were provided granulated sugar, powdered milk, and water ad libitum For all assays, adult house flies (3-5 d old) we re aspirated from the screened rearing cages using a handheld vacuum with a modified crevice tool attachment Flies used for the laboratory assays were placed into a 5 C environment for 5 min to subdue act ivity. Flies were then placed on a chilled aluminum tray, sexed, and counted. H ouse flies used for field cage assays were not

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31 anesthetized, but were as pirated from the screened rearing cage s and released directly into field cages. Laboratory Arenas. Arenas (31 x 25 x 21 cm) were constructed using PVC pipe (1.27 cm [0.5 in]) (Figure 3-1A). Rubber bands were us ed to establish individual treatment positions; four treatment positions were us ed in the cord attractiveness bi oassay and five positions were used in the impregnated cord bioassay. All co rds were attached to the treatment positions vertically using paper clips and were uniformly di stributed along the length of the arena. The cord attractiveness bioassay held two randomly a ssigned cords at each tr eatment position and the impregnated cord bioassay held only one cord at each treatment position according to a 5 x 5 Latin square configuration. Aren as were enclosed with a transp arent plastic bag (3716 cm2 [24 x 24 in], 1 mil poly, Uline, Waukegan, IL). Cord Attractiveness Bioassay. Eight cords were evaluated: nylon (Braided, MultiPurpose Braid 75 lb. load limit, Welli ngton Cordage LLC, Madison, GA), polypropylene (Braided, Multi-Purpose Rope 56 lb. load limit, Wellington Cordage LLC, Madison, GA), cotton (Braided, Multi-Purpose Sash Cord 28 lb. load limit, Wellington Cordage LLC, Madison, GA), cotton wick (Sterilized roll #200209, Richmond Dental Company, Charlotte, NC), manila (Twisted, Natural Rope 108 l b. load limit, Wellington Cordage LLC, Madison, GA), wool (Twisted, Natural Cord, Wooded Haml et Designs, Greencastle, PA), leather (Tan laces, #6192, Rothco, Ronkonkoma, NY), and parachut e cord (550 test, white, purchased locally from M & C Army Surplus Store, Gainesville, FL). Fifty female flies were released into each arenas and 10% sugar water was provided ad libitum Number of flies resting on cords was co unted every 10 min for 2 hr. Arenas were lightly shaken between each count to displace flies from their res ting positions. Four replications

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32 were performed in the laboratory (28 1C) under continuous light on separate days using different flies. Impregnated-Cord Laboratory Bioassays. Cotton, manila, wool, polypropylene, and nylon cords were selected from the cord attrac tiveness experiments to be evaluated in the impregnated-cord experiments. Each impregnatedcord experiment consisted of 6 arenas. Five arenas were organized into a 5 x 5 Latin s quare design, blocking for treatment position, and a sixth arena was used as a control. The control arena had no treated cords and all cords within it maintained the same cord positions throughout all experiments (left to right: position 1 = cotton; 2 = wool; 3 = manila; 4 = polypropylene; 5 = nylon). Separate experiments were done to evaluate cords (15.24 cm length [6 in], 0.6 cm [0.25] diam) impregnated with a 0.1% fipronil or a 0.6% indoxacarb solution. The 0.1% fipronil solution was prepared by combining 2.7 ml of the formulated insecticide (Termidor SC, 9.1% a.i., BASF, Research Triangle Park, NC) with 250 ml of ta p water. The 0.6% indoxacarb solution was prepared by combining 5 g of formulated insecticide (DPX MP062, 30WG, DuPont, Wilmington, DE) with 250 ml of tap water. Cords were impregnated by dipping for ~2 sec in the insecticide solution and were then allowed to dry in a fume hood. Groups of 50 female flies were placed within each arena and provided a 10% sugar water solution ad libitum Mortality counts were recorded until at least 80% mortality was observed. Due to the differences in the mode of action of the insecticides, mortality for flies exposed to fipronil-impregnated cords was defined as the in ability to remain standing; flies exposed to indoxacarb-impregnated cords were considered dead if they were unresponsive to touch. Each experiment was run in the laboratory (28 1C) under continuous light and replicated twice.

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33 Impregnated-Cord Field Cage Bioassay. Cages (1.8 x 3.7 x 1.8 m) were constructed from PVC pipe (2.54 cm [1 in] diam) and en closed with mesh screening (Outdoor Cage, #1412A, 18 x 14 mesh, Bioquip, Rancho Dominguez, CA). Black plastic sh eeting (6 mil) was used to line the floor. A sampling stage, cons tructed of two vertical cinder blocks and an inverted storage bin (Palletote #1721, 37 liter, Rubbermaid, Winchester, VA), was placed in the center of the cage (Figure 3-1B). On top of the sampling stage there were two 994-ml (1 qt) chick waterers, one filled with 10% sugar water and the other with tap water, and a 60-ml plastic cup filled with 8 g of previously used larval house fly medium. The chick waterers provided enough sustenance for the duration of the test and the plastic cup was used as an attractant. The plastic cup was covered with a paper towel and s ealed with a rubber band to prevent flies from ovipositing on the medium. Treatments consisted of two long (0.9 m) and eight short (0.6 m) lengths of 0.1% fiproniland 1.2% indoxacarb-impregnated wool cords. The 0.1% fipronil solution was prepared by combining 7.7 ml of the formulated insectic ide (Termidor SC, 9.1% a.i., BASF, Research Triangle Park, NC) with 700 ml of tap water. The 1.2% indoxacarb solution was prepared by combining 28 g of formulated insecticide (D PX MP062, 30WG, DuPont, Wilmington, DE) with 700 ml of tap water. Each cord was treated in the same manner as the laboratory experiments, except the cords were dipped and soak ed for 1 min prior to drying. Depending on fly availability, 27.5 35 ml (9.8 1.8 flies/ml) of flies was released into each cage. After a 1-h acclimation period, pretreatment fly counts were taken. Before fly counts were taken, the operator wa lked three laps around the interior of the cage to disturb flies from their resting positions and to recover any dead flies from the cage floor. Four consecutive fly counts were then taken from the outside of the cage 1 min after exiting. All flies th at landed

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34 on the sampling stage, chick waterers, and plastic cup attractant were counted. Treatments were then hung vertically from the mesh ceiling using pa per clips in specific locations (Figure 3-1C) and post-treatment fly counts were taken at 24 a nd 48 h using the same method described above. After the initial 48 h evaluation, tr eatments were aged in the elements for four weeks, at which point residual effectiveness was re-evaluated as described above. Three replicates were performed at each treatment age (0 and 4 wk). Statistical Analysis. All statistical analyses were performed using JMP IN (SAS Institute 2005), except probit analysis estimates were performed using SAS (S AS Institute 2001). For the cord attractiveness experiments, the mean number of flies/cord wa s analyzed using a one-way analysis of variance and contrasts were performed between natural and synthetic cords and the animaland plantbased cords. For the laboratory insecticide-im pregnated cord laboratory experiments, mortality data were corrected using Abbo tts formula (1925) and arcsine s quare root-transformed. A twoway analysis of variance was performed on the 24h fipronil data and the 48-h indoxacarb data to determine if treatment position ha d an effect on mortality. LT50 values were estimated by probitanalysis regression (Finney 1971). Potency ratios, using the cotton cord as the standard, were performed using the method described in R obertson and Preisler (1991). Slopes, LT50 values, and potency ratios were consider ed significantly different if the 95% confidence intervals did not overlap. For the field cage experiments, percen t fly count reductions were calculated from the control fly counts. Fly count reductions and mortal ity data (number of dead flies recovered from cage floor) were then analyzed for each treatment age (0 and 4 wk). A ll means were separated using the Students T or St udent-Newman-Keuls test ( = 0.05).

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35 Results In the laboratory studies, all flies fully recovered from chilling after approximately 45 min at which point the flies were disp ersed throughout the entire arena. Flies were more attracted to the manila cord, which had significantly more flies resting on it than any ot her cord (Figure 3-2). No significant differences were seen between the other natural cords or between the synthetic cords; however, all synthetic cords had significan tly less flies resting on them than the natural cords (F: 112.69, df = 368, P = <0.001) and the plantbased cords were more attractive than the animal-based cords (F: 11.64, df = 368, P = <0.001) The least attractive cord was the nylon parachute cord. The laboratory design had no position or interacti on effects for either fi pronil (F: 1.05; df = 4; P = 0.3982, F: 0.8347; df = 20; P = 0.6583) or indoxacarb (F: 0.71; df = 4; P = 0.5906, F: 0.32; df = 20; P = 0.9955) in the insecticide-impr egnated cord experiments. At the 24 h (fipronil) and 48 h (indoxacarb) recordings, all impregnated-cords had significantly higher mortality than the controls (Figure 3-3). House flies suffered significantly higher mo rtality when exposed to the fipronilimpregnated wool cord than any other fipronilimpregnated cord at 24 h (93%). The other fipronil-impregnated natural cord s had percent mortalities below 15%, with manila causing only 5% mortality at 24 h. No significant differences in mortality were seen between the fipronilimpregnated nylon and polypropylene cords or the fipronil-impregnated cotton and manila cords at 24 h. The indoxacarb-impregnated wool cord cause d significantly higher mo rtality (85%) than any of the other cords except for the cotton cord at the 48 h record ing. No significant differences in mortality were seen between the syntheti c indoxacarb-impregnated cords or between the cotton and manila indoxacarb-impregnated cords. Si gnificant differences in mortality were seen

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36 between the wool and manila indoxacarb-impr egnated cords. The indoxacarb-impregnated nylon cord caused the lowest mortality at 48 h (47%). Fiproniland indoxacarb-impregnated cords effi cacy results can be viewed in Table 3-1. In general, the fipronil impregnated cords had lower LT50 and LT90 values than the indoxacarbimpregnated cords. Among the fipronil-impregna ted cords, the wool cord had the lowest LT50 and LT90 values and the impregnated co tton cord had the highest LT50 and LT90 values. The LT50 values for the synthetic cords were relatively low compared to the other cords, but the LT90 values were no different from the cotton cord. The manila cord LT50 value was the second highest, but had the second lowest LT90 value behind the wool cord; it is important to note that it also had the highest slope compared to the other cords. All cords were more effective than the cotton cord except for the nylon cords LT90 value. All indoxacarb-impregnated cords had LT50 values >32 h and LT90 values >51 h. The indoxacarb-impregnated wool cord had lower LT50 and LT90 values than all other indoxacarbimpregnated cords except for the manila cord, which showed no significant differences in LT50 values. The indoxacarb-impregnated polypr opylene and cotton cords each had LT50 values of 52 h and LT90 values >100 h, which were the highest values for the experiments. No differences in LT values were observed between the manila and nylon cords. All cords were more effective than the cotton cord except for the polypropylene cords LT50 and LT90 values. In the field cage experiments, no significant differences in fly count reductions occurred between the treatments (Table 3-2). Both trea tments had >57% fly count reductions by 24 h and >87% by 48 h, independent of the treatment age. Dead flies were collected from all cages at every recording; significantly more were collected from the treatment cages than the controls. Fipronil treatments had significantly more dead flies than the indoxacarb treatments with fresh

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37 cords at 24 and 48 h and with aged cords at 24 h. No significant differenc es were seen between the number of dead flies collected from the fipron il treatments and indoxacarb treatments at the 4 wk, 48-h recording. Discussion Insecticide-impregnated fly cords are base d on a fundamental component in a flys behavior flies prefer to rest on objects with distinct edges, such as twigs, wires, cord, and line (Scudder 1949). Since insecticide-impregnate d cords only represent a small proportion of available resting surfaces availa ble to flies, it is assumed th at factors which enhance a flys attraction to the cords would be be neficial to the effectiveness of the treatment. Surfaces which are more attractive to flies would be expected to cause quicker mortality because of increased exposure to the insecticide. Arevad (1965) f ound flies to favor dark, rough surfaces over light, smooth surfaces. Specific factors influencing a flys attraction to natural fiber cords were evaluated by Fay and Lindquist (1954 ). They found sisal cord to be more attractive than jute or wool cords of the same size, but less attractive than a similar sized cott on cord. When given a choice between only cotton and sisal cords, flies pr eferred the sisal cord. They also found that the same type of cord was more attractive to flies as the cord diameter increased between 0.131.1 cm. In our attractiveness experime nt, the cords we evaluated vari ed by fiber type (animal or plant), color, texture, and, in some cases, even diameter. All of the natural cords we evaluated were more attractive than the synthetic cords. The natural cords were rougher than the relatively smooth synthetic cords; in addition, the plant-fibered manila cord and the animalfibered leather cord were darker than the other cords. These factors may have increased their overall attractiveness to the flies. If compari ng the most attractive cord (manila) to the least attractive (parachute cord) the differences in te xture and color are subs tantial (Figure 3-4).

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38 Manila is a very rough, coarsely textured brown thatch cord made from the leaf fibers of the abaca tree, Musa textiles, while the parachute cord is a rela tively smooth kernmantle cord made of white nylon. The parachute cord was one of two cords less than 0.64 cm, which may have decreased its attractiveness. The other cord less than 0.64 was the leat her cord, but it was as attractive as the other natural cord s (except manila) despite its diam eter being half the size. The leather cords dark color may have increased its attractiveness or it may have been more attractive due to animal odors that were still associated with the material. The previously available commercial fly cord s were exclusively made of cotton. Cotton was cheap, durable, absorbent and widely availabl e. Although cotton was relatively attractive in our experiments, it had very poor efficacy for bot h fipronil and indoxacarb when compared to the other natural and synthetic cords te sted indicating that it may not be the best type of cord to use for insecticide treatment. Fiproniland i ndoxacarb-impregnated wool cords had the greatest efficacy in our experiments despite flies resting on it 50% less than the manila cord in the cord attractiveness experiments. This is contrary to the previous assumption that quicker mortality would result from increased exposure to a more attractive insecticide-impregnated cord and neglects to account for the inse cticide-substrate inte raction. Highly organic materials readily bind to pesticides and make them less effectiv e (Dell et al. 1994, Gardner et al. 2000) and may have accounted for the low LT50 and LT90 values seen in the cotton cords and in the LT50 value of the fipronil-impregnated manila cord. The ex act reason wool outperfor med the other cords in our experiments was not fully i nvestigated, but it is likely due to the insecticide-substrate interaction. The wool cord was the only anim al-fibered cord evalua ted and is naturally impregnated with several oils. Both fipronil a nd indoxacarb are very lipop hilic insecticides and

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39 probably dissolved readily within these oils which likely increased the rate of insecticide transfer from the cords through the waxy layers of the flys cuticle. In the laboratory, the indoxacarb cords genera lly provided a much slower kill than the fipronil cords, however, in the field cages differen ces were not as apparent. Indoxacarb is a proinsecticide that needs to be bioactivated within th e insect before it is toxic and will always cause mortality slower than an insecticide, such as fi pronil, that is toxic upon contact once a lethal dose is obtained. Flies poisoned by indoxacarb in the laboratory are shielded from desiccation and predation, which may have proved to be vital to their prolonged survival in the laboratory experiments. Furthermore, the indoxacarb dose wa s increased in the field cage experiments and may have affected the faster results seen in th e field cage experiments. Both treatments showed a decrease in efficacy in the field experiments afte r being aged 4 weeks, but still had adequate fly count reductions and causing significantly more flies to die than the control. In conclusion, the use of inse cticide-impregnated cords is very practical to supplement a house fly management program. Insecticide-im pregnated cords ensure adequate residual coverage in areas difficult to trea t with traditional residual insect icides and they can easily be removed and relocated to other fl y resting areas if needed or a lternated with cords impregnated with other active ingredients to reduce the possibi lity of resistance development. More research still needs to be done to determ ine adequate doses and rates of treatment, keeping in mind that these may vary depending on cord type and insec ticide used. Wool cord outperformed all other cords evaluated in this study and fipronil and indox acarb both appear to be effective insecticides for house fly control.

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40Table 3-1. Efficacy of various cords impregnated with 0.1% fipronil or 0.6% indoxacarb on female house flies. Lethal Times (h) (95% CL) Potency Ratio (95% CL) Treatment Cord n Slope SE 50 90 2 P 50 90 Fipronil Cotton 2750 9.52 0.36b 39.7 (39.2-40.2)e 54.1 (52.9 55.5)c 9.060 0.1067 1.00 e 1.00 d Manila 1000 12.98 0.83a 35.0 (34.5-35.6)d 44.0 (42.6 45.7)b 1.213 0.5445 1.13 (1.12-1.15)d 1.23 (1.21-1.25)b Wool 1250 5.32 0.29c 12.9 (12.3-13.4)a 22.4 (21.3 23.8)a 0.650 0.4200 3.09 (3.01-3.16)a 2.42 (2.35-2.48)a Polypro 2500 4.65 0.29d 26.2 (25.6-27.0)c 49.6 (46.4 53.8)c 3.643 0.7249 1.51 (1.45-1.57)c 1.09 (1.04-1.14)c Nylon 1500 3.68 0.25e 23.0 (21.2-24.6)b 51.3 (48.2 55.4)c 1.455 0.6927 1.72 (1.66-1.79)b 1.05 (1.00-1.11)cd Indoxacarb Cotton 2248 4.04 0.13c 52.2 (50.3-54.3)c 108.5 (102.1-115.9)c 1.3494 0.5093 1.00 c 1.00 c Manila 1659 5.10 0.74b 36.2 (32.3-38.7)ab 64.7 (60.2 73.4)b 3.4030 0.3336 1.44 (1.35-1.54)ab 1.68 (1.54-1.82)b Wool 4250 6.44 0.16a 32.6 (32.0-33.1)a 51.5 (50.1 53.1)a 8.6295 0.2804 1.60 (1.54-1.67)a 2.11 (2.01-2.20)a Polypro 2250 3.11 0.20d 52.2 (49.7-54.5)c 134.8 (122.7-151.7)d 0.6717 0.7147 1.00 (0.91-1.10)c 0.80 (0.72-0.90)c Nylon 2000 6.57 0.54a 39.2 (37.2-40.8)b 61.5 (59.4 64.4)b 2.9321 0.2308 1.33 (1.27-1.39)b 1.76 (1.68-1.85)b Total number of trials; 500 flies/trial except for the cotton (498) and manila (487) indoxacarb-impregnated cords (Probit [SAS Institute 2002]). Mortality was corrected using Abbotts Formula. Means within a column, in the same treatment group, followed by the same lett er are not significantly different based on non-overlap of 95% confidence intervals.

PAGE 41

41 Table 3-2. Cumulative number of dead flies and percent fly count reduction in relation to control fl y counts of house flies ex posed to 0.1% fiproniland 1.2% indoxacarbimpregnated cords in field cages. % Fly Count Reduction SEM # of Dead Flies Treatment Age Treatment 24 h 48 h 24 h 48 h 0 Weeks Fipronil 80.22 13.10a 98.66 1.34a 83.0 1.0a 95.3 3.4a Indoxacarb 57.39 6.92a 97.21 1.43a 30.3 4.4b 59.7 2.0b Control 4.7 4.2c 11.3 8.5c 4 Weeks Fipronil 59.25 22.10a 87.43 12.57a 53.0 7.8a 79.3 7.2a Indoxacarb 64.39 5.89a 87.72 7.34a 30.3 4.7b 62.0 8.1a Control 3.3 1.7c 10.3 2.0b Cumulative mean number of f lies recovered from cage floor. Means in a column, within the same treatment age, followed by the same letter are not significantly different (P > 0.05; Stud ents T or Student-Newman-Keuls test)

PAGE 42

42 Figure 3-1. Laboratory and fiel d experimental design elemen ts. A). Laboratory arena constructed of PVC pipe. Cords were suspended between the rubber bands using paper clips. B) Sampling stage used in fi eld cage experiments. Chick feeders with either 10% sugar water or tap water and a plastic cup containing previously used larval medium was used as sustenance and attraction. C). Cord placement in relation to sampling stage (SS) in the fi eld cage bioassay. Crossed circles were short cords (0.6 m) and empty circles were long cords (0.9 m).

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43 Figure 3-2. Attraction of female house flies to various natural and synthetic cords.

PAGE 44

44 Figure 3-3. Female house fly mortality exposed to various natural and synthetic cords treated with 0.1% fipronil for 24 h (A) a nd 0.6% indoxacarb for 48 h (B).

PAGE 45

45 Figure 3-4. Comparison of the most attractive co rd (manila) and the least attractive cord (nylon parachute) in the cord attr activeness experiments.

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46 CHAPTER 4 EVALUATION OF A NEW IMIDACLOPRID BAIT FOR HOUSE FLY CONTROL Introduction The house fly, Musca domestica L., is the most commonly encountered pest of the generalized group of Diptera called filth flies. Large numbers of house flies are frequently found in areas where manure, garbage, and other decaying organic matter are abundant. Although primarily a nuisance to people and animals, house flies can pose a health risk by mechanically transferring pathogens picked up fr om their breeding sites, particularly when they enter homes or eating establishments. When the source of infest ation is inaccessible or sanitation measures are not effective, house fly control is often achieve d using dry insecticidal scatter baits. Two widely used scat ter baits are Maxforce Granular fly bait and Golden Malrin fly bait. Maxforce Granular is an imidacloprid-based bait cont aining the fly attractant (Z)-9-tricozene, the bittering agent Bitrex, and other attractants and inert ingredients. It is currently the only imidacloprid-based scatter bait available. Golden Malrin contains 1.1 % methomyl, 0.049 % (Z)-9-tricosene, as well as other attractants and in ert ingredients, and is one of several methomylbased scatter bait formulations available. In general, dry scatter baits have many adva ntages over other types of insecticidal fly control products: they are easier to work with in field environments, they can be more attractive to flies than liquid baits, and they usually ha ve a longer storage shelf life (Gahan et al. 1954, Darbro and Mullens 2004). However, dry scatter baits need to be replaced frequently in some areas when granules become covered by manure or other debr is (Barson 1987). The U.S. Environmental Protection Agency (EPA), acting under legislative mandates, also requires the scatter bait granules be dyed to distinguish th em from other non-toxic materials. For example, the Maxforce Granular fly bait is formulated as red granules and the Golden Malrin is

PAGE 47

47 formulated as blue granules. When these granules become wet, the dye often bleeds onto the surrounding surface and may be unsightly for the user. Label restrictions are also very different and can limit the uses of certain active ingredients or insecticide products. Golden Malrin can only be applied as scatter bait or within bait stations, whereas, the Maxforce Granular can be applied as scatte r bait, within bait stations, or it can be mixed with water and painted onto surfaces allowing it to be applie d directly to distinct fly resting areas, such as on ceilings or rafters. However the use of Maxforce Granular is more restricted than that of Golden Malrin because its label restricts its use in food establishments. Golden Malrin, despite being the only carbamate-based in secticide not classified as restricteduse, can be used within food establishments when used in bait stations placed at least 1.2 m from the ground in areas where food proces sing or preparation does not occur. An imidacloprid sprayable bait, Maxforce Fly Spot, has recently become commercially available. It contains 10% imid acloprid, 0.1% Z-9-tr icosene, Bitrex, and inert ingredients. This formulation still maintains the advantages of trad itional scatter baits, while eliminating some of the disadvantages of the currently ava ilable products. Once applied, Maxforce Fly Spot bait dries clear and the label allows fo r application within food establishm ents when the facility is not in operation. Our objectives were to compare the effectiven ess of the new sprayable bait in relation to the two most commonly used dry scatter baits. In addition, we compared the performance of the imidacloprid sprayable and granul ar baits in a controlled fiel d environment and tested the imidacloprid sprayable bait im pregnated in cords.

PAGE 48

48 Materials and Methods Insects. The Horse Teaching Unit (HTU ) strain of house flies, M. domestica L., reared at the University of Florida in Gainesville was used for all experiments. Larvae were reared on a diet medium, modified from Hogsette (1992), containing 3 liters wheat bran, 15 ml methyl paraben, 1.5 liters water, and approximately 200 g (250 ml) dairy calf feed (Calf Manna pellets, Manna Pro Corp., St. Louis, MO). A ll developmental stages were held at 26 1 C and 55% RH with a 12:12 (L:D) photoperi od. Adult flies emerged within in screened rearing cages and were provided granulated s ugar, powdered milk, and water ad libitum For all assays, adult house flies (3-5 d old) we re aspirated from the screened rearing cages using a handheld vacuum with a modified crevice tool attachment Flies used for the laboratory assays were placed into a 5 C environment for 5 min to subdue act ivity. Flies were then placed on a chilled aluminum tray, sexed, and counted. H ouse flies used for field cage assays were not subdued, but were aspirated from the screened re aring cages and released directly into field cages. Laboratory Arena Design. Arenas (31 x 25 x 21 cm) were constructed using PVC pipe (1.27 cm [1/2 in] diam) (Figure 3-1A). Rubber bands were used to establish five uniformly distributed cord positions along the length of the arena. Each position held one cord which was vertically attached to the rubbe r bands using paper clips. Fi ve cords were used with all laboratory experiments: nylon (Braided, Mult i-Purpose Braid 75 lb. load limit, Wellington Cordage LLC, Madison, GA), polypropylene (Braid ed, Multi-Purpose Rope 56 lb. load limit, Wellington Cordage LLC, Madison, GA), cotton (B raided, Multi-Purpose Sash Cord 28 lb. load limit, Wellington Cordage LLC, Madison, GA), manila (Twisted, Natural Rope 108 lb. load limit, Wellington Cordage LLC, Madison, GA), and wool (Twisted, Natural Cord, Wooded

PAGE 49

49 Hamlet Designs, Greencastle, PA). Arenas were enclosed with a transparent plastic bag (3716 cm2 [24 x 24 in], 1 mil poly, Uline, Waukegan, IL). Field Cage Design. Cages (1.8 x 3.7 x 1.8 m) were c onstructed from PVC pipe (2.54 cm [1 in] diam) and enclosed with mesh scr eening (Outdoor Cage, #1412A, 18 x 14 mesh, Bioquip, Rancho Dominguez, CA). Black plastic sheeting (6 mil) was used to line the floor. A sampling stage, constructed of two verti cal cinder blocks a nd an inverted storage bin (Palletote #1721, 37 liter, Rubbermaid, Winchester, VA), was placed in th e center of the cage (Figure 3-1B). On top of the sampling stage there were two 994-ml (1 qt) chick waterers, one filled with 10% sugar water and the other with tap wate r, and a 60-ml plastic cup filled with 8 g of previously used larval house fly medium. The chick waterers pr ovided enough sustenance for the duration of the test and the plastic cup was used as an attractant. The plastic c up was covered with a paper towel and sealed with a rubber band to prev ent flies from ovipositing on the medium. Fly Bait Comparisons. Three fly baits, 2 dry scatter ba its and 1 sprayable bait, were applied to polystyrene Petri dishes (100 by 15 mm; Fisher Scientific, Pittsburgh, PA). The methomyl granular bait (Golden Malrin, Methomyl 1.1%, (Z)-9-Tricosene 0.049%, Wellmark International, Schaumburg, Illinois; dose: 0.23 g/0.9 m2) and the imidacloprid granular bait (Maxforce Granular fly bait, Bayer CropS cience, Kansas City, MO; dose: 30.17 g/0.9 m2) were sprinkled on the Petri dish. The Imidaclopr id sprayable bait (Maxforce Fly Spot bait, Imidacloprid WG 10, Lab Code: 342/207-7, Bayer CropScience, Monheim am Rhein, Germany; dose: 0.45 g/0.9 m2; rate: 0.12 g Pr/ml/0.9 m2) was suspended in tap water, sprayed on the Petri dish bottom using an airbrush (Paasche, Type H, Chicago, IL), and allowed to dry in a fume hood prior to being placed in the arena. Bait di shes were placed on the bottom rubber bands in the center of the arena. A separate arena with an untreated Petri dish was used as the control.

PAGE 50

50 Cords in these experiments served only as resting positions for the flies, they were untreated and hung in the same configuration for all repetitions: (left to right: pos ition 1 = cotton; 2 = wool; 3 = manila; 4 = polypropylene; 5 = nylon). Groups of 50 female flies were placed within each arena and a 10% sugar water solution was provided ad libitum Mortality was recorded at 1, 3, 5, and 24 h. Flies were considered dead if they were unable to stand or fly. Each experiment was run in the laboratory (30 1C) under continuous light and replicated three times. The two imidacloprid baits were evaluated in the field cages. Treatments consisted of two plastic lattice squares (0.19 m2) treated with imidacloprid granul ar bait, imidacloprid sprayable bait, or tap water (control). Trea tments were applied on only one si de of the lattice at the same rates as the laboratory fly bait comparison assa ys. The imidacloprid granular bait was mixed with tap water (1.44 g: 1 ml) and pa inted on. The tap water (3.78 ml/0.9 m2) and imidacloprid sprayable bait (0.12 g Pr/ml/0.9 m2) were sprayed on using an airb rush. All treatments were allowed to thoroughly dry outdoors in the open air before being hung on the ceiling PVC pipes using cable ties. Each lattice square was pl aced medially along the length of the cage, approximately 0.5 m away from each side of th e sampling stage and positioned so that the treated surfaces of th e lattice squares faced opposite directions. Depending on fly availability, 27.5 35 ml (9.8 1.8 flies/ml) of flies were released into each cage. After a 1-h acclimation period, pretreatment fly counts were taken. Before fly counts were taken, the operator wa lked three laps around the interior of the cage to disturb flies from their resting positions and to recover any dead flies from the cage floor. Four consecutive fly counts were then taken from the outside of the cage 1 min after exiting. All flies th at landed on the sampling stage, chick waterers, and plastic cup attractant were counted. Treatments were

PAGE 51

51 then hung within the cages and post-treatment fly counts were taken at 1 and 24 h using the same method described above. After th e initial 24 h evaluation, treatments were aged in the elements for two weeks, at which point residual effectiv eness was re-evaluated as described above. Three replicates were performed at ea ch treatment age (0 and 2 wk). Bait-Treated Cords. Five laboratory arenas were orga nized into a 5 x 5 Latin square design, blocking for treatment position, and a sixt h arena was used as the control. Each treatment consisted of a cord (15.2 cm length, 0. 6 cm diam) impregnated with a 2.5% solution of imidacloprid sprayable bait. The imidacloprid solution was prepared by combining 25 g of the formulated insecticide with 100 ml of tap wate r. Cords were impregnated by dipping for ~2 sec in the insecticide solution and were then allowed to dry on aluminum foil covered trays in a fume hood prior to being placed into the arenas. The control arena had no treated cords and had the same cord configuration as the fly bait comparison bioassay described above. Laboratory tests were conducted wi th groups of 60 female flies/arena. Flies were provided a 10% sugar water solution ad libitum Morbidity (knockdown) was recorded at 2-5 h posttreatment and mortality was recorded at 24, 48, and 72 h. Flies were considered knocked down if they did not move when touched at the 2-5 h recordings. Flies that were unresponsive to touch at the 24, 48, and 72 h recordings were consid ered dead. Each experiment was run under continuous light in the same laboratory conditio ns as described above and replicated twice. In the field cages, treatments consisted of tw o long (0.9 m) and eight short (0.6 m) lengths of imidacloprid-impregnated wool cords. Each cord was treated in the same manner as the laboratory experiments, except the cords were dippe d and soaked for 1 min prior to drying. Flies were released and fly counts were taken in the same manner as the imidacloprid bait field cage experiments. Cords were hung vertically from the mesh ceiling using paper clips in specific

PAGE 52

52 locations, which remained constant throughout the experiment (Figure 3-1C). Post-treatment sampling counts were done at 24 and 48 hrs. Af ter the initial 48 h eval uation, treatments were aged in the elements for four weeks, at which point residual effectiveness was re-evaluated as described above. Three replicates were performed for each treatment age (0 and 4 wk). Data Analysis. All analyses were done using a one-way analysis of variance with JMP IN (SAS Institute 2005). For the fly bait compar ison and the bait-treated cord experiments, percent morbidity (bait-treated cords) and mortal ity data were arcsine square root-transformed and analyzed for each time interval. For the fi eld cage experiments, percent fly count reductions were calculated from the control fly counts. Fly count reductions and mortality data (number of dead flies recovered from cage floor) were then analyzed for each treatment age (0 and 2 wk for the imidacloprid comparisons; 0 and 4 wk for th e bait-treated cord e xperiments) (Conover and Iman 1981). Means for all analyses were separa ted using the Students T test or the Student Newman Kuels (SNK) method ( = 0.05). Results In all laboratory experiments, flies did not fully recover from chilling until roughly 1 h after entry into the arenas. Flies first contacte d the baited Petri dishes in the bait comparison experiments approximately 35 min post recovery in the following order: imidacloprid granular bait, imidacloprid sprayable bait, methomyl granular bait. Initial fly contact on the treated cords in the bait-treated cord e xperiments was not observed. Fly Bait Comparisons. The imidacloprid granular and th e imidacloprid sprayable baits had higher fly mortality than the methomyl gran ular fly bait at 3 h, but by 24 h the methomyl granular bait had the highest ove rall mortality (Figur e 4-1). At 24 h, fly mortality with the imidacloprid sprayable bait was not significantly different from morta lity with either the imidacloprid granular or the methomyl granular fly baits, but signifi cant difference in fly

PAGE 53

53 mortality did exist between the methomyl and the im idacloprid granular fly baits. All treatments were significantly different than the c ontrol fly mortality at all observations 3 h. In the imidacloprid field cage experiments, no differences were seen between either treatments with fresh or aged cords (Table 4-1) Both treatments had >35% fly count reductions at 24 h and >70% fly count reduc tions by 48 h with fresh cords, but fly count reductions did not exceed 8% with aged cords for either treatmen t. A fly count increase was observed with the imidacloprid granular treatment at 48 h with aged cords. The number of dead flies collected in the treatment cages was significantly different from the control with fresh cords, but no differences were observed between the aged treat ments and controls 2 wk post-treatment. Bait-Treated Cords. Morbidity increased on a time-depe ndent basis until approximately 3-4 hours post-treatment, at which time flies re covered from being knocked down by all of the imidacloprid bait-treated cords ex cept for the cotton cord (Figure 4-2). Flies exposed to all imidacloprid bait-treated cords had knockdown recovery by 24 h. All cords caused significantly more mortality than the control cords at ev ery 24 h recording (Figure 4-3). Beyond 24 h, mortality with the imidacloprid ba it-treated nylon, cotton, and wool cords increased more sharply than the mortality caused by the bait-treated ma nila and polypropylene cords. The imidacloprid bait-treated wool cord caused th e highest overall fly mortality (74%). All other cords resulted in house fly mortalities <60% with the imidacloprid ba it-treated polypropylene cord showing the lowest overall fly mortality (25%). In the field cages, the imidacloprid bait-trea ted cords caused >87% fly count reductions by 24 h with fresh and aged cords (Table 4-2). Th e aged bait-treated cord s fly count reductions decreased by ~6% by 48 h, whereas fly count reductions in creased by ~6% with the fresh cords.

PAGE 54

54 The number of dead flies collected was significantly different than the control at 24 and 48 h for both fresh and aged cords, except for the 48 h recording with the aged cords. Discussion When insecticides are used for house fly cont rol, most users expect to see satisfactory results (i.e. dead flies) within hours and markedly reduced populations within 1-2 days. Thus, an effective fly bait will attract flies quickly and cause high mortality within a relatively short period of time. In our bait comparison experiment flies contacted the im idacloprid baits sooner than they contacted the methomyl bait, which ma y have been a contributi ng factor to the higher fly mortality at 3 h with the imidacloprid baits th an with the methomyl bait. However, the higher fly mortality with the methomyl bait after 24 h suggests that methomyl may be a more potent, although slower acting, active ingredient. Other studies comparing imidacloprid and me thomyl baits have also shown the same mortality trends we observed in flies exposed to technical and ba it formulations of imidacloprid and methomyl (White et al. 2007). In those experi ments, White et al. observed up to 50% of the flies that were knocked down by imidacloprid form ulations recovered. They hypothesized innate characteristics, independent of resistance m echanisms, may make some flies tolerant to neonicotinoids. We observed knockdown recovery in the bait-treated cord experiments with all cords, but no recovery was seen in the house flie s exposed to any of the baits we tested in the bait comparison experiments. Recovery may have occurred in these experiments, but was not observed because recordings were not taken between the 5 h and 24 h recordings. Flies that were knocked down were not isolated from the arena in our experiments and could have received a second dosing before having the oppor tunity to fully recover. Differences in cord material or treatment app lication technique may have also attributed to fly recovery in the bait-treated cord experiments. Cord satu ration is dependent on the cord

PAGE 55

55 composition and may have lead to sublethal dosi ng. We observed that cord composition varied between the types of cords we used, and even am ong individual cords. Distribution of oils and other materials on the surface of each cord make it difficult to have the bait uniformly distributed over the surface of the cord. When bait is spraye d onto a solid surface, such as the Petri dish, a precise amount of bait remains on the surface after the water evaporates. However, when a cord is dipped into a bait solution, the bait may be absorbed deep into the fibers, disperse throughout, or pool in areas on the cord. Thus, some bait may not be available for flies to contact. This is evident when hand-dipping cords in dyed insectic ides materials, such as in an indoxacarb wettable granule (WG) solution, which is gray ish-brown in color. Despite being fully submerged in the insecticide solutions, some of the cord often remains its natural color, apparently void of any bait. Once the same cords are allowed to completely dry and are removed from the drying trays, brown staining surrounds we re they once lay indicating that some of the insecticide may be lost during the drying process as well. It is undetermined if knockdown re covery occurred in our field cage experiments. If flies are knocked down in the field, natural enemies may prey upon them before they are able to fully recover. We observed knocked-down flies bein g preyed upon by ants, spiders, and lizards. Others, undoubtedly, became victim to desicca tion after being knocked down. Barson (1987) found that flies knocked down by methomyl in the fiel d often lost their ability to fly, but still had the ability to reproduce. White et al. (2007) commented that 10% of the flies knocked-down by imidacloprid in their laboratory studies fully recovered and resumed normal behavior when protected from a second exposure to imidacloprid. The inability to fly would make flies more vulnerable to predation by natural enemies. Howeve r, if reproduction is stil l occurring, it will be detrimental to any fly control program because of a flys prolific reproductive capabilities. Field

PAGE 56

56 studies to determine the effect of knock down and recovery on a fly management program would be beneficial. Insecticide-impregnated cords have been used extensively in th e past to control flies and have recently been examined using new insecticid es not yet registered fo r use against house flies ( Unpublished, Chapter 3). In those experiments, indox acarb and fipronil were more effective on wool cords than any other natural and synthetic cords tested. Wool co rds also showed higher efficacy than the other natural and synthetic co rds tested when treated with the imidacloprid sprayable bait. Exact LT50s were not determined in these experiments because of the knockdown and subsequent recovery observed, but based on Figure 4-3, we estimate that 50% of the flies died after approximately 60 hours (2.5 d) with the wool cord. With such a long period to reach 50% fly mortality, imidacloprid-treate d cords were slower acting than any of the fipronilor indoxacarb-impregnated cords previously tested. However, in the field cages, the imidacloprid bait-treated cords reduced fly counts by 80% in 24 h. The high lipid content of wool cord may facilitate the tr ansfer of insecticide through the cuticle of the house fly, but this does not explain the differences in results be tween the laboratory a nd field assessments. Differences in the rate of cords per cage area ma y explain the differences in the laboratory and field results. The cords in the field cages were hung at a rate of 9.1 m of cord/9.3 m2 area based on the recommended rate of the insecticide cords used in the 1950s (Fehn 1958, Smith 1958, Weinburgh et al. 1961). The rate in the laborat ory arena was comparatively much lower, 0.02 m of cord/m2 vs. 1.0 m of cord/m2. This rate appears to be quite high and may vary between different cord/insecticide combinations. A dditionally, the aforementioned predation and desiccation of the knocked down f lies probably was significant fact or contributing to the rapid fly count reductions in the field cages.

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57 When comparing the imidacloprid bait-treate d cords and the imidacloprid bait-treated lattice squares, the bait-t reated cords were more effective. The lattice squares did not reduce the fly counts to any significant degree after being aged 2 weeks, but the imidacloprid bait-treated wool cords had good fly count redu ctions even after be ing aged 4 weeks. The plastic lattice squares were selected as a treatment surface to represent the material found on many portable toilets, latrines, or dumpsters. Bait treatments on this type of surface are very vulnerable to environmental conditions because the material does not allow the bait to penetrate as in the cord treatments. Damp conditions in the morni ngs and unexpected preci pitation (6.5 cm) that occurred between evaluation intervals washed aw ay most of the bait product from the lattice squares. With the imidacloprid granular bait applic ation, the lattice squares were almost completely void of the red dye following these mo isture events and red staining was seen on the cage floor. We assume that the imidacloprid sp rayable bait was also washed off the lattice squares given the results of th e fly count reductions and the d ead flies recovered, but was not observed because the bait has no color. The ba it-treated cords were exposed to 3.5 cm less precipitation than the bait-treated lattice squares, which may have also aff ected the bait available on the cord. When dipping cords in a bait soluti on, the bait is absorbed in between individual cord fibers and even deeper into the core of the cord, making the bait more protected from environmental conditions. When th e cords are then subjected to th ese moisture events, the bait may concentrate in specific areas of the cord (suc h as the cord end) instead of completely leaving the cord as seen with the lattice. Additionally, flies prefer to rest on cords and probably receive a larger dose of insecticide in this manner as compared to when they land on the flat surfaces of the lattice. When a fly lands on a flat surface they are exposed only to the precise amount of

PAGE 58

58 toxicant that absorbs through thei r tarsi or is imbibed; however when resting on cords, their thorax and abdomen are also brushed by the treated cord fibers. In conclusion, the imidacloprid sprayable bait was found to be as effective as the traditional commercial scatterbaits compared in this study. Its unique formulation and less restrictive product label allow this bait to be used in areas where other fly baits are prohibited. Unless a more rain-fast formulation becomes availa ble, the imidacloprid sprayable bait will need to be reapplied frequently in areas with high mois ture or precipitation especially when applied to non-absorbent surfaces such as portable latrin es or dumpster lids. The baits potential effectiveness in insecticide-impregnated cord s needs further investigation due to differing laboratory and field results. Regardless, this ne w imidacloprid sprayable bait should prove to be a very useful tool in any fly management program.

PAGE 59

59Table 4-1. Number of dead a nd percent fly count reduction in relation to control fly counts of house flies exposed to imidaclo prid bait-treated lattice s quares in field cages. % Fly Count Reduction SEM # of Dead Flies Treatment Age Treatment 1 h 24 h 1 h 24 h 0 Weeks Imidacloprid granular bait 47.1 6.3a 70.9 4.4a 36.0 10.0a 117.0 9.5a Imidacloprid sprayable bait 36.6 20.5a 80.2 4.7a 36.3 2.0a 113.0 10.1a Control 0.3 0.3b 1.7 0.7b 2 Weeks Imidacloprid granular bait 0.8 8.2a -3.8 20.1a 1.0 0.6a 19.7 11.2a Imidacloprid sprayable bait 7.6 10.8a 6.3 19.5a 0.7 0.7a 14.0 11.6a Mean number of individuals recovered from cage floor. Means in a column, within the same treatment age, followed by the same letter are not significantly different (P > 0.05; Stud ents T test or Student-Newman Kuels Method)

PAGE 60

60 Table 4-2. Number of dead and percent fly count reduction in relation to control fly co unts of house flies exposed to imidacloprid bait-treated cords in field cages. % Fly Count Reduction SEM # of Dead Flies Treatment Age Treatment 24 h 48 h 24 h 48 h 0 Weeks Bait-Treated Cords 90.4 4.1 96.8 3.2 97.7 17.2a 114.3 19.8a Control 12.7 4.6b 19.7 7.7b 4 Weeks Bait-Treated Cords 87.9 0.9 82.4 14.7 50.7 14.0a 69.3 23.7a Control 8.7 3.8b 13.0 4.7a Mean number of individuals recovered fr om cage floor. Means in a column, within the same treatment age, followed by the same letter are not significantly diffe rent (P > 0.05; Students T test)

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61 0 10 20 30 40 50 60 70 80 90 100 0510152025Time (h)% Mortality Methomyl Granular Imidacloprid Granular Imidacloprid Sprayable Control Figure 4-1. Mortality of female house flies e xposed to imidacloprid and methomyl granular scatter baits and a spraya ble imidacloprid bait.

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62 0 2 4 6 8 10 12 14 16 18 20 012345Time ( h ) % Morbidit y Cotton (plant) Manila (plant) Wool (animal) Nylon (synthetic) Polypropylene (synthetic) Control Figure 4-2. Morbidity (knockdown) of female h ouse flies exposed to natural and synthetic cords dipped in a 2.5% solution of imidacloprid sprayable bait.

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63 0 10 20 30 40 50 60 70 80 90 100 0244872 Time (h)% Mortality Cotton (plant) Manila (plant) Wool (animal) Nylon (synthetic) Polypropylene (synthetic) Control Figure 4-3. Mortality of female house flies exposed to natural and synthetic cords dipped in a 2.5% solution of imidacloprid sprayable bait.

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64 CHAPTER 5 SUMMARY AND CONCLUSIONS House flies are often found in tremendous numbers in situations where U.S. troops are most frequently deployed, such as areas stressed by conflict or natural disasters. House flies are nuisance pests that decreases troop morale and they pose a health risk to deployed troops that may disrupt mission objectives if diseases that are associated with their pr esence stress the health care systems. The main objective for this research was to evaluate insecticide-impregnated cords and sprayable fly bait as new methods to control the house fly and provide usable information to the DOD that will help protect the deployed war fighter. First and foremost, house flies can be contro lled using insecticide-impregnated cords and sprayable fly bait and their use would benefit agricultural, urban, and military fly management programs. Insecticide-impregnated cords and sp rayable fly baits are bo th very easy to use products. Impregnated cords can be hung usi ng a staple gun or other similar method and sprayable bait is mixed with wa ter and sprayed onto any surface. Impregnated cords can be removed and relocated quickly, which may be bene ficial in relatively mobile troop deployments or in situations were resistance is suspected. Sprayable baits can be removed with simple water wash down and easily reapplied when needed. One of the most interesting findings in th is research is the further understanding in insecticide-impregnated cord toxicity. Previ ous insecticide-impregnated cords were made exclusively of cotton because it was cheap, durable, and relative ly attractive to house flies, however, the cotton cords in our experiments we re the least effective cord. Wool cords consistently showed they were more effective th an any of the other cord s we evaluated despite the fact that they were not the most attractive co rd to the house flies. House flies were most attracted to the highly organic manila cord and pref erred to rest on it more than any other natural

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65 or synthetic cord evaluated. The wool cord wa s the only animal-based cord impregnated with insecticides and possibly gave it a distinct adva ntage over the other cords because it is naturally coated with oil, which probably helped facilitate inse cticide transfer thro ugh the fly cuticle. Two fly scatter baits, Maxforce Granular and Golden Malrin fly bait, are both listed on the DOD pesticide contingency list for house fly cont rol but both have limitations. Only one can be applied in food service areas and both can stain materials and equipment which can compromise camouflage and substrate appearan ce. Eliminating these disadvantages, while maintaining bait efficacy, would appear to provide advantages to a fly management program and benefit the deployed war fighter. The ne w sprayable imidacloprid bait, Maxforce Fly Spot, is as effective as the two previously mentioned scatter baits in the la boratory and as effective as its counterpart, Maxforce Granular, in the field. Its unique formulation allows it to be used within food serving areas and it will go unnoticed because it dries clear. Unfortunately, like the other bait products, environmental factors, such as ra in, decrease the efficacy over time. Residual efficacy did improve when the bait was applied to the cords rather than the non-absorbent plastic surfaces often found on latrines and dumpsters. We anticipated that the research completed here would provide some information that could be further used to develop future products that could be benefit the DOD. The insecticides evaluated in the impregnated cord studies are both non-registered for house fly control. House flies have shown little to no resistance towa rds fipronil and indoxacarb and both insecticides appear to be very effective agai nst this pest. At this time, no information has been obtained on whether or not the manufacturers of fipronil or indoxacarb are seeking registrations for these products to control flies. Howe ver, a Colorado company has inform ed me of their interest in indoxacarb-impregnated cords and has begun conversations with DuPont regarding further

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66 research into this type of product. The spraya ble imidacloprid fly bait is not currently listed on the DOD pesticide contingency list but it rece ived its EPA registration in 2006 and became commercially available in mid 2007. A formal re quest will be submitted to the Armed Forces Pest Management Board this summer to request that it be assigned a National Stock Number (NSN) and be placed on the DOD pesticide c ontingency list. If successful, Maxforce Fly Spot will be readily available to the deployed war fighter for use in their fly management programs.

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67 APPENDIX A REVIEW OF INSECTICID E-IMPREGNATED CORDS The use of insecticide-impregnated cords to co ntrol house flies was first tried with DDT in 1947 (Baker et al.). By the early 1950s, im pregnated materials for fly control became increasingly common (Pimentel et al. 1951). Ins ecticide-impregnated cords were being used on dairies, at rural residences, military mess halls, state fairs, and state prisons with great success (Kilpatrick 1955, Maier and Mathis 1955, Sor oker 1955, Kilpatrick and Schoof 1956). Commercial Fly-Cords distributed by Fly-Cord Inc. (Savannah, Ge orgia) were widely available and used by 1957 (Fehn 1958, Smith 1958). These cords were considered the treatment of choice for use in buildings housing animals because of the economy and efficiency (Fay and Kilpatrick 1958). Fay and Lindquist (1954) recogn ized that impregnated cords offer only a small percentage of the surfaces availabl e for flies to rest. Exploiting the f actors which enhance a flys attraction to a particular cord would subsequently lead to higher mortality on impregnated cords. They found that cord type, thickness, and color significantly influenced a flys attr action to a particular cord. Sisal and cotton cords were more attractive than jute or wool cords. Cord attractiveness increased with cord diameters between 3/64 and 7/16. Flies preferred red and black cords over blue, yellow, green, or white cords. No pr eference was evident between vertically or horizontally hung cords. Although several other organophosphate insectic ides have been evaluated for their effectiveness, all provided satisfactory resu lts. However, only one cord was available commercially (Kilpatrick and Schoof 1959, Gratz et al. 1964, Rabari and Patel 1976). The commercial Fly-Cord was a 3/32 diameter, re d cotton cord impregnate d with 13.79% parathion and 3.54% diazinon (Fehn 1958, Smith 1958). The co rd was supplied on a reel containing 300

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68 feet; each linear foot of cord contained 75-100 mg of parathion (Youngblood 1960). The manufacturer recommended a rate of 30 linear feet of Fly-Cord per 100 square feet of floor area in locations were adult flies congregate (F ehn 1958, Smith 1958). Cords were normally hung about three feet apart using staple guns or simply tied on to the structure (Fehn 1958, Youngblood 1960). If multiple competing resting su rfaces were available to the flies, most applicators increased the amount of impregnated co rds in the area. In places where flies were not seen resting or where condi tions were unsuitable (i.e. area s with drafts), no cords were placed. The use of impregnated-cords was an attempt to find new methods to reverse the resistance associated with residual spraying of chlorina ted hyrdrocarbons such as DDT (Keiding and Jespersen 1986). Fly-Cords offered an easyto-use control method that restricted and concentrated a residual insectic ide. This method lowered the se lection pressure for resistance because flies which avoided contact with the co rds diluted the remaining population of resistant individuals (Keiding and Jespersen 1986). Other methods, such as paint-on baits, non-residual space sprays, and combining baits and larvicides, were evaluated and determined effective (Keiding and Jespersen 1986). Pain t-on baits and non-residual space sprays are widely available and used today in the United States. No commercially available insecticide-impre gnated cord products are currently available in the United States. This is partially becau se the main active ingredients, parathion and diazinon, are not registered for filth fly control. In addition, the use of selective inse cticidal baits has become increasingly popular. In Denmar k, the use of impregnated cords was abandoned because of newer construction techniques that allo wed for more ventilation in the animal shelters and more effective residual sprays became ava ilable (Keiding and Jespersen 1986). The World

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69 Health Organization and the United States Military continue to r ecommend the use of insecticide treated or impregnated cord s (Rozendaal 1997, AFPMB 2006)

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70 APPENDIX B REVIEW OF FLY BAITS Insecticidal baits have long been used for fly c ontrol. One of the original bait formulations contained either 1-2% formaldehyde or sodium ar senite mixed with milk or sugar water (Keiding 1976). Residual insecticides have been mixed with sugar to make them more attractive to flies and serve as a type of ba it, but this type of mixt ure is often not as effective as baits specifically formulated to attract flies. Todays fly baits are loaded with many diff erent attractants including pheromones, sugars, and other substances that spec ifically attract house flie s. Most of these fly baits are formulated as either dry scatter baits, but many also come in easy-to-use bait station devices. Some of the dry scat ter baits can be mixed with wa ter and painted on a surface. Insecticidal baits have many advantages over other chemical control methods. Baits are relatively inexpensive, usually have a longer storag e shelf life, are more attractive to flies than other chemical control methods, and are easier to work with in field environments (Gahan et al. 1954, Darbro and Mullens 2004). The dry scatter ba its simply get scattered on the ground in the infested areas or placed within a bait station while bait station de vices normally only need to be opened and hung in infested locations. Keiding ( 1976) considered baits le ss likely to select for resistance than residual sprays. He was most likely referring to the phys iological resistance seen in many of the organophosphate and carbamate insect icides at that time. Today, many insects, including flies, have been shown to develop be havioral resistance to ba its (Darbro and Mullens 2004). Fly baits do have disadvantages. Dry scatter baits do not target fly resting areas unless they can be painted on and this type of application often leads to stained surfaces because the U.S. Environmental Protection Agency (EPA) requi res the scatter bait gra nules to be dyed to distinguish them from other non-toxic materials. Staining can also occu r when these granules

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71 become wet by rain and bleed onto the surroundi ng surface, which may be considered unsightly for the user. Bait station devices are also degrade rapidly in e nvironments with intense sun or precipitation. Dry scatter baits also need to be replaced frequently in some areas when granules become covered by manure or other debr is such as garbage (Barson 1987). As with any chemical insecticide, label restrictions can be disadvantageous (although necessary in most cases) and limit the needs of the applicator. For example, the two most widely used dry scatter baits in the United States are Maxforce Granular fly bait and Golden Malrin fly bait. Maxforce Granular is an imidacloprid-based ba it (neonicotinoid class) containing the fly attractant (Z)-9-tricozene, the bittering agent Bitrex, and other attractants and inert ingredients. Golden Malrin contains 1.1 % methomyl (carbamate class), 0.049 % (Z)-9tricosene, as well as other attractants and inert in gredients, and is one of several methomyl-based scatter baits available. Maxforce Granular has a more restricted label than that of Golden Malrin because its label restricts its use in food establishments. Golden Malrin, despite being the only carbamate-based insecticide not classified as a restricted-use insecticide, can be used within food establishments when used in bait stations placed at leas t 1.2 m from the ground in areas where food processing or pr eparation does not occur. Recently, a new fly bait has become commercia lly available that may offer advantages over some of the other current fly baits available. Maxforce Fly Spot bait contains 10% imidacloprid, 0.1% Z-9-tricosene, Bitrex, and inert ingredients. Once applied, Maxforce Fly Spot bait dries clear and the label allows for a pplication within agricu ltural livestock production facilities and serving areas of food establishments when the facility is not in operation.

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72 APPENDIX C REVIEW OF INSECTICIDES EVALUATED Fipronil Fipronil is a phenylpyrazole insecticide that wa s first registered in the United States in 1996 (Connelly 2001). It is used to control termites, an ts, roaches, fleas, ticks, and various other agriculture and turf pests. No fipronil products are currently re gistered for house fly control. Fipronil causes mortality by cont act and ingestion (Vargas et al. 2005). Insects exposed to fipronil show extreme neural excitation that eventu ally leads to insect pa ralysis and death. Death is caused by the disruption of the normal passage of chloride ions through the -aminobutyric acid type A (GABA) receptor system of insects (Scharf et al. 2000). Hainzl et al. showed fipronil to have a tighter binding affinity toward insect GABA-re gulated chloride channels over mammalian receptors (1998). Fipronil-sulfone, an important active metabolite of fipronil, was also found to block the glutamate receptors in co ckroaches (Zhao et al. 2004). Glutamate-gated chloride channels are only found in invertebrate systems at skeletal neuromuscular junctions of both the peripheral and central nervous system (Raymond and Sattelle 2002, Scharf 2003). The unique quality of fipronil to affect two target sites makes it a highly selective insecticide and potentially important factor limiti ng the development of detectable resistance (Zhao et al. 2004). To date, resistance to fipronil a ppears to remain at low levels or even be non-existent in house flies (Scott and Wen 1997, Scott et al. 2000, Kris tensen et al. 2004). Low levels of crossresistance have been reported in multi-resi stant house flies and at tributed to monoxygenasemediated detoxification, decrease d insecticide penetra tion, and target site mutations (Wen and Scott 1999, Liu and Yue 2000, Kristensen et al. 2004). Resistance surveys in New York found house flies susceptible to fipronil even at LC99 levels (Scott et al. 2000).

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73 Indoxacarb Indoxacarb (DPX-MP062) is an oxdiazine insectic ide that was first regi stered in the United States in 2000 (EPA 2000). It is a 75:25 mixture of the active S-isomer (DPX-KN128) and the inactive R-isomer (DPX-KN127). A less effective formulation (DPX-JW062) contains a 50:50 mixture of the two stereoisomers. Indoxacarb was originally formulated to control lepidopteran pests of fruits and vegetables, but newer registrations include cockroach, mole cricket, and fire ant baits. It is not curren tly registered for house flies. Indoxacarb is considered an organophosphate replacement and designated as a reducedrisk insecticide by the EPA. It is a pro-insecticide that must be biochemically converted to a toxic decarbo-methoxyllated metabolite (Dias 2006). Toxicological effects are dependent on the conversion of the inactive metabolite to its toxi c form within the insect body. In mammals, indoxacarb metabolites are rapidly excreted; whereas in insects, indoxacarb is rapidly converted by an esterase and amidase into DCJW, which is the more insecticidally active metabolite. Insects exposed to lethal doses of indoxacar b experience impaired nerve function, feeding cessation, paralysis, and eventually death. Indoxacarb poisoning occurs through contact or ingestion and it works by blocking the sodium cha nnel of the insect nervous system. This mode of action is distinct from other insecticides th at target the sodium ch annels of insects (DDT, pyrethroids) because DCJW disrupts the sodium channels without modifying the activation or deactivation kinetics (Lapied et al. 2001). It works by blocking the channel pore, and prevents normal sodium ion flow. Because indoxacarb is a new chemistry, not much work has been done on insecticide resistance. Shono et al. (2004) se lected house flies that had >118-fo ld resistance in as little as three generations and concluded that the resistance mechanism was associated with a major

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74 factor on autosome 4 and a minor factor located on autosome 3, both of which are not linked to any resistance mechanisms previously described. Imidacloprid Imidacloprid is a chloronicotinyl nitroguanidine (neonicotinoid) that wa s first registered in the United States in 1994 (NPTN 1998). It is us ed to control a wide va riety of agricultural, urban, public health, and veterinary pests and is estimated to a ccount for 11-15% of the total global insecticide market (Tomizawa and Casida 2005). Several formulations are available for different treatment applications, only two ar e available for house fly control: Maxforce Granular fly bait and Maxforce Fly Spot bait. Both products are baits, but differ from one another by their formulation. Maxforce Granular fly bait is a red gr anule that can be applied as a traditional scatter bait, within a bait stati on, or mixed with water and painted onto a surface. The Maxforce Fly Spot bait, alternatively, is white we ttable powder that is mixed with water and sprayed onto a surface. When the Maxforce Granular fly bait is pain ted on a surface, or if the granules become wet, it stains the surface red, whereas the Maxforce Fly Spot bait is clear and does not stain. Imidacloprid kills insects through contact and ingestion by agonizing the nicotinic acetylcholine receptor (nAChR) (Fossen 2006). In house flies, imidacloprid is metabolized by oxidation to the olefin metabolite, which has th e same toxicological activity as imidacloprid (Nishiwaki et al. 2004). Insects exposed to le thal doses of imidacloprid experience nervous system excitability, modified f eeding behavior, and death. Imidacl oprid is considered a selective insecticide because: (1 ) imidacloprid has a higher affinity for the insect nAChRs than mammalian nAChRs, and (2) there are more nAChR s located in the insect nervous system than what are found in mammalian systems (Yamamoto et al. 1995).

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75 Resistance has yet to be reporte d in house flies (Gao et al. 2007), but it is well-documented in Drosophila (Daborn et al. 2001). Cross resi stance has been observed in multi-resistant house flies (Wen and Scott 1997). Multiple resistan ce mechanisms are suspected in house flies. Monoxygenase-mediated detoxication seems to be a primary mechanism in some strains of house flies, but not in others (Wen and Scott 1997, Liu and Yue 2000). Methomyl Methomyl is a carbamate insecticide that was first registered in the United States in 1968 (EPA 1998). It is used to control a wide vari ety of agricultural, ur ban, public health, and veterinary pests. Several methomyl formulations are available, but the 1% fly bait formulation is the only one which is not classified as a restricted-use pesticide. Methomyl causes mortality by contact and inge stion by inhibiting the acetylcholinesterase (AChE) enzyme, which occurs in the central ne rvous system. Methomyl binds to AChE and prevents it from binding to acetylcholine. This re sults in acetylcholine sa turation at its neural receptor, which results in a dramatic increase in generation of nerve impulses. Insects exposed to methomyl show signs of hyperexcitabi lity, convulsions, paralysis, and death. Decarbamoylation of AChE is rapid and, therefor e, carbamates are considered reversible AChE inhibitors and recovery fr om sub-lethal poisonings can occur quickly (Yu 2007). Little resistance to methomyl has been seen in house flies despite its frequent use and the high levels of resistance seen in house flies to other carbamat es (Barson 1989, Webb et al. 1989, Scott et al. 2000, Darbro and Mullens 2004). F lies feeding on methomyl granules have been found to receive a super-lethal dose that may play a large roll in why resistance has not been as widespread (Price and Chapman 198 7). Behavioral resistance, or bait aversion, has started to become more apparent. In 1989, Barson (1989) re ported that 8% of the resistant flies were repelled by methomyl. In a study comparing the mortality of 35 field strains of house flies fed

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76 methomyl in choice and no-choice tests, mortality decreased by nearly 30 % when the flies were given the choice test (D arbro and Mullens 2004).

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77 LIST OF REFERENCES Abbot, W. S. 1925. A method of computing the effectiv eness of an insecticide. J. Econ. Entomol. 18. AFPMB. 2006. Filth flies: significance, surveillance and control in contingency operations, pp. 1-59. In G. B. White [ed.], TG 30. Armed For ces Pest Management Board, Washington, D.C. Anderson, J. R. 1964. The behavior and ecology of various flies associated with poultry ranches in northern California, pp. 30-34, 32nd Ann. Conf. Calif. Mosq. Assn. Inc. Proc. and Pap. Arevad, K. 1964. On the orientation of houseflies to va rious surfaces. Entomol. Exp. Appl. 8: 175-188. Axtel, R. C. 1999. Poultry integrated pest management: status and future. J. Poultry 4: 53-73. Baker, W. C., H. I. Scudder, and E. L. Guy. 1947. The control of houseflies by DDT sprays. Pub. Med. Health Rep. 62: 597-612. Barson, G. 1987. Laboratory assessment of different met hods of applying a commercial granular bait formulation of methomyl to control adult houseflies ( Musca domestica L. ) in intensive animal units. Pestic. Sci. 19: 167-177. Barson, G. 1989. Response of insecticide-resistant and susceptible houseflies ( Musca domestica ) to a commercial granular bait form ulation containing methomyl. Med. Vet. Entomol. 3: 29-34. Beck, A. F., and J. E. C. Turner. 1985. A comparison of five house-fly (Diptera: Muscidae) population monitoring techniques. J. Med. Entomol. 22: 346-348. Bishopp, F. C., and L. W. Laake. 1921. Dispersion of flies by fli ght. J. Agric. Res 21: 729-766. Brookes, V. J. 1956. The nutrition of the larva of the house fly, Musca domestica L. (Muscidae: Diptera), pp. 114, Entomology. Uni v. of IL, Champaign-Urbana. Buchan, P. B., and R. S. Sohal. 1981. Effect of temperature and di fferent sex ratios on physical activity and life span in the adult housefly, Musca domestica Exp. Geront. 16: 223-228. Burrus, R. G. 2005. Humanitarian efforts following the Indonesian tsunami. Vector Ecol. Newsletter 36: 11-12. Connelly, P. 2001. Environmental fate of fipronil, pp. 1-17. Calif. Environ. Prot. Agency, Sacramento. Conover, W. J., and R. L. Iman. 1981. Rank transformations as a bridge between parametric and nonparametric statistics. The American Statistician 35: 124-129.

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78 Daborn, P., S. Boundy, J. Yen, B. Pittendrigh, and R. ffrench-Constant. 2001. DDT resistance in Drosphila correlates with Cyp6g1 over-expression and confers crossresistance to the neonicotinoid imidacl oprid. Mol. Genet. Genomics 266: 556-563. Darbro, J. M., and B. A. Mullens. 2004. Assessing insecticide resistance and aversion to methomyl-treated toxic baits in Musca domestica L (Diptera: Muscid ae) populations in southern California. Pest Manag. Sci. 60: 901-908. Dell, C. J., C. S. Throssell, M. Bischoff, and R. F. Turco. 1994. Estimation of sorption coefficients for fungicides in soil and tu rfgrass thatch. J. En viron. Qual. 23: 92-96. Dias, J. L. 2006. Environmental fate of indoxacarb, pp. 20. Depart. Pest. Reg., Sacramento. EPA. 1998. Methomyl, pp. 8, R.E.D. Facts. Environmental Protection Agency. EPA. 2000. Indoxacarb fact sheet, pp. 17. Environmental Protection Agency. Fay, R. W., and D. A. Lindquist. 1954. Laboratory studies on factor s influencing the efficiency of insecticide impregnated cords for house fl y control. J. Econ. Entomol. 47: 975-980. Fay, R. W., and J. W. Kilpatrick. 1958. Insecticides for control of adult Diptera. Annu. Rev. Entomol. 3: 401-420. Fehn, C. F. 1958. House fly control at Scout camps with insecticide-impregnated cords. Calif. Vect. Views 5: 62-63. Finney, D. J. 1971. Probit analysis. University Press, Cambridge. Fossen, M. 2006. Environmental Fate of Imidacloprid, pp. 16. Depart. Pestic. Reg., Sacramento. Gahan, J. B., H. G. Wilson, and W. C. McDuffie. 1954. Dry sugar baits for the control of houseflies. Agrie Food Chem 2: 425-428. Gao, J. R., J. M. Deacutis, and J. G. Scott. 2007. The nicotinic actylcholine receptor subunit Md6 from Musca domestica is diversified via post-trans criptional modification. Insect Mol. Biol. 16: 325-334. Gardner, D. S., B. E. Branham, and D. W. Lickfeldt. 2000. Effect of turfgrass on soil mobility and dissipation of cyproconazole. Crop Sci. 40: 1333-1339. Geden, C. J. 1995. Natural enemies of house flies, pp. 77-82. In H. H. J. Van Horn [ed.], Nuisance concerns in animal manure manage ment: odors and flies. Florida Cooperative Extension, Gainesville, FL. Geden, C. J. 2005. Methods for monitoring outdoo r populations of house flies, Musca domestica L. (Diptera: Muscidae). J. of Vector Ecol. 30: 244-250.

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79 Georghiou, G. P., and A. Lagunes-Tejeda. 1991. The occurrence of resist ance to pesticides in arthropods: An index of cases reported th rough 1989. Food and Agriculture Organization of the United Nations. Graczyk, T. K., B. H. Grimes, R. Knight, A. J. Da Silva, N. J. Pieniazek, and D. A. Veal. 2003. Detection of Cryptosporidium parvum and Giardia lamblia carried by synanthropic flied by combined fluorescent in situ hybridization and a mo nclonal antibody. Am. J. Trop. Med. Hyg. 68: 228-232. Gratz, N. G., G. Sacca, and J. Keiding. 1964. Dichlorvos for the control of houseflies. Riv. Parasite 25: 269-278. Greenberg, B. 1960. House fly nutrition. II. comparative surv ival values of sucrose and water. Ann. Entomol. Soc. Am. 53. Greenberg, B. 1973. Flies and disease: biology and diseas e transmission. Princeton University Press, New Jersey. Grubel, 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: 1300-1303. Hainzl, D., L. M. Cole, and J. E. Casida. 1998. Mechanisms for selective toxicity of fipronil insecticide and its sulfone me tabolite and deulfinyl photoproduct. Chem. Res. Toxicol. 11: 1529-1535. Hecht, O., R. Muni z, and A. Nava. 1968. Contrary responses of Musca domestica concerning their selection of different shades and hues. Ent. Exp. Appl. 11: 1-14. Hogsette, J. A. 1992. New diets for production of house f lies and stable flies (Diptera: Muscidae) in the laboratory. J. Econ. Entomol. 85: 2291-2294. Hogsette, J. A. 1995. The house fly: basic bi ology and ecology, pp. 71-76. In H. H. J. Van Horn [ed.], Nuisance Concerns in Animal Manur e Management: Odors and Flies. Florida Cooperative Extension, Gainesville, FL. Howard, J. 2001. Nuisance flies around a la ndfill: patterns of abundan ce and distribution. Waste Manage. Res. 19: 308-313. Howard, J. J., and R. Wall. 1998. Effects of contrast on attraction of the housefly, Musca domestica to visual targets. Med. Vet. Entomol. 12. Jones, C. J., S. A. Isard, and M. R. Cortinas. 1999. Dispersal of synanthr opic Diptera: lessons from the past and technology for the fu ture. Ann. Entomol. Soc. Am. 92: 829-839.

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85 BIOGRAPHICAL SKETCH Jeffrey Conrad Hertz was born in 1976 in Peoria, Illinois. His parents, Marion Conrad Butch Hertz and Margaret El oise Weezie Hertz (Hilton), raised him in Lewistown, Illinois. They moved to Bernadotte, Illinois where he continued to attend school in neighboring Lewistown until he graduated from Lewistown Community High School in 1994. He entered the United States Navy and reported to basic training at Re cruit Training Command, Great Lakes, Illinois in November later that same year. Over the last 12 years, he served with the United States Marine Corps, at Naval hosp itals, and most recently, he was assigned to the medical staff at the United States Capitol. He received his Associate of Science degree in medical laboratory technology from George Wa shington University, Washington D.C. in 2002 and his Bachelor of Science in interdisciplinary studies, majo ring in biology from Mountain State University, Beckley, West Virginia in 2003. In 2004, he wa s selected as the very first enlisted Sailor selected to study entomology under the Medical Service Corps In-service Procurement Program (MSC-IPP). Upon gra duation HM1 (FMF) Jeffrey Hertz will be commissioned to the rank of Li eutenant Junior Grade as a medi cal entomologist in the Medical Service Corps. He enjoys running and is an acti ve member of Centennial Lodge #174 of Ancient Free and Accepted Masons locate d in Upper Marlboro, Maryland. He, his wife, Karina, and two children, Conrad and Kyra, are excited about th eir upcoming move to Jacksonville, Florida where he will be working at the Navy Entomological Center of Excellence.