The evaluation of poultry pest management techniques in Florida poultry houses

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Title:
The evaluation of poultry pest management techniques in Florida poultry houses
Uncontrolled:
Poultry pest management techniques in Florida poultry houses
Physical Description:
xvii, 308 leaves : ill. ; 28 cm.
Language:
English
Creator:
Hogsette, J. A ( Jerome Adkins ), 1945-
Publication Date:

Subjects

Subjects / Keywords:
Poultry -- Diseases -- Control   ( lcsh )
Housefly -- Control   ( lcsh )
Mites -- Control   ( lcsh )
Poultry industry -- Health aspects -- Florida   ( lcsh )
Genre:
bibliography   ( marcgt )
theses   ( marcgt )
non-fiction   ( marcgt )

Notes

Thesis:
Thesis--University of Florida.
Bibliography:
Includes bibliographical references (leaves 267-290).
Statement of Responsibility:
by Jerome Adkins Hogsette, Jr.
General Note:
Typescript.
General Note:
Vita.

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Source Institution:
University of Florida
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All applicable rights reserved by the source institution and holding location.
Resource Identifier:
aleph - 000014212
notis - AAB7411
oclc - 06300526
System ID:
AA00003479:00001

Full Text









THE EVALUATION OF POULTRY PEST MANAGEMENT
TECHNIQUES IN FLORIDA POULTRY HOUSES











BY

JEROME ADKINS HOGSETTE, JR.






















A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL
OF THE UNIVERSITY OF FLORIDA IN
PARTIAL FULF!LLiENTT OF THE REQUIREMENTS FOR
THE DEGREE OF DOCTOR OF PHILOSOPHY


UNIVERSITY OF FLORIDA


1979












ACKNOWLEDGEMENTS

I would like to express my gratitude to Dr. Jerry F. Butler for

his guidance, advice, and encouragement while serving as the Chairman

of the Supervisory Committee. Thanks are also extended to Drs. P. G.

Koehler and D. W. Hall for serving on the Supervisory Committee and

aiding in the completion of the dissertation.

Much appreciation is extended to Drs. R. A. Voitle and C. R.

Douglas for serving on the Supervisory Committee and for their

suggestions and constructive criticism during the course of this

research.

I am extremely grateful to Dr. R. H. Harms for allowing me to

utilize the facilities at the University of Florida Poultry Science

Department. Additional thanks are expressed to.Dr. R. B. Christmas

and his farm crew for their cooperation and assistance during the

mite trials at Chipley, Florida.

Myriad thanks are extended to Diana Simon and the many laboratory

technicians and fellow graduate students who participated in various

phases of this research.

And finally, warm thanks are extended to my wife, Debbie, for

her perseverance, patience, and understanding; and for the wonderful

sense of humor she maintained while typing this dissertation.












TABLE OF CONTENTS

PAGE

ACKNOWLEDGEMENTS................................................. i

LIST OF TABLES.......................................... .......... vi

LIST OF FIGURES................................................... xii

ABSTRACT................................................. ........ xv

INTRODUCTION....................................................... 1

LITERATURE REVIEW.................................................. 3

The House Fly................................................... 3
History and Economic Importance............................... 3
Bionom ics................................................. ... 4
Methods for Larval Control................................. .. 7
Methods for Adult Control in Poultry Houses................... 21
Northern Fowl Mites............................................. 27
Description, Biology, and Control ............................ 27
Effects on the Host...................... ...... ...... .......... 32
Medical Importance of Northern Fowl Mites..................... 34
Control of Northern Fowl Mites................................ 36

METHODS AND MATERIALS.............................. .............. 43

Laboratory Trials............................................... 43
Environmentally Controlled Rearing Conditions................. 43
Colonization and Rearing of Flies............................ 43
Dissection and Mounting of Cephaloskeletons of
Third-lnstar Fly Larvae........................... ..... .... 47
Bioassay of Poultry Manure ................................... 48
Addition of a Liquid Insect Growth Regulator (IGR)
to Larval Media of Flies............................ ........ 49
Laboratory Tests with Granular Baits.......................... 49
Topical Application of Insecticides to House Fly Adults....... 50
Laboratory Bioassay of Acaricides............................ 51
Field Trials.................................................... 52
Rotovation.................................................... 52
Description of the Tilling Site............................... 54
Monitoring Larval Fly Populations............................. 54
Poultry and Poultry Facilities used when Evaluating
IGR's as Oral Larvicides.................................... 57
Calculation of Hen-Day Production and Average Daily
Feed Consumpton ................................................. 60
Addition of IGR's to Poultry Feed............................ 61







PAGE


Topical Application of Granular IGR's to Poultry Manure....... 61
Mixing and Application of Liquid IGR's and
Organophosphorus Larvicides................................. 63
Addition of a Liquid IGR to the Drinking Water of Hens......... 63
Placement of Light Traps ..................................... 63
Field Tests with Granular Baits................ .............. 64
Application of Contact Residuals to Selected Surfaces.......... 67
Application of Contact Residuals to Plywood Panels............. 67
Evaluation of Northern Fowl Mite Populations.................. 69
Field Application of Acaricides to Caged Hens................. 71
Field Application of Acaricides to Floor Birds................ 72
Compounds Utilized for Fly or Mite Control ................... 72
Treatment of Data............................................. 72

RESULTS.......................................................... 76

House Flies..................................................... 76
Manure Management ............................................. 76
C-7.; .': aenescens Basic Biology Studies .................. ..... 91
Competition Studies with /Ermetia ll-ucns.................... 11
insect Growth Regulators and Organophosphorus Larvicides...... 123
Blacklight Electrocutor Grid Traps for Adult Fly Surveys...... 161
Efficacies of Granular Fly Baits.............................. 167
Contact Residuals............................................. 183
Northern Fowl Mites ............................................. 19
Dosage-Mortality Curves for Selected Acaricides............... 194
Control of Endemic Florida Strains of Northern Fowl
Mites with Carbaryl, Malathion, and Ravap................... 199
Efficacy of Two Synthetic Pyrethroid Compounds Against
Northern Fowl Mites on Laying Hens in Floor Pens............. 206
The Effects of Northern Fowl Mites on Egg Production.......... 212

DISCUSSION........................................................ 233

The Value of Rotovation as a Method of Manure Management......... 239
Op:.,r aenescens Larvae as Predators of 2usca domestic
Larvae........................................................ 247
Morphological Proof that (Ophyra aenescens is Predaceous .......... 248
The Value of Ophyra aenescens as a Biocontrol Agent............. 249
Rearing Ophyra aenescens in the Laboratory...................... 249
The Influence of Larvae of Revmetia iZucens on Other
Species of Fly Larvae......................................... 250
The Efficacy of Dimilin as a Feed Additive...................... 251
The Efficacy of Methoprene as a Feed Additive................... 252
Methoprene as a Topical Larvicide............................... 232
Laboratory Studies with CGA 72662............................... 253
CGA 72662, Dimethoate, Dichlorvos, and Ravap as
Topically Applied Larvicides............. ............. ...... 253
The Efficacy of CGA 72662 in Water.............................. 254







PAGE

Light traps for Surveying Adult Fly Populations................. 255
Granular Baits for House Fly Control............................ 255
Efficacy of Synthetic Pyrethroids as Contact Residuals.......... 257
Susceptibility of Endemic Florida Strains of Northern Fowl Mites
to Carbaryl, Malathion, Ravap, and Synthetic Pyrethroids...... 258
The Effects of Northern Fowl Mites on Egg Production............ 260
Evaluation of the Mite Rating System............................ 262

CONCLUSIONS....................................................... 264

LITERATURE CITED................................................... 267

APPENDICES

1A RAW DATA FROM FIRST OPHYRA AZNTZCE' ADULT LONGEVITY
STUDY.................................................... 292
1B RAW DATA FROM SECOND OPHYRA ,5'TSJ:'X ADULT LONGEVITY
STUDY... ................................................. 293
!C RAW DATA FROM THIRD OPHYRA AENESCENS ADULT LONGEVITY
STUDY.................................................... 294
1D RAW DATA FROM FOURTH OPHYIR AENESCENS ADULT LONGEVITY
STUDY.................................................... 296
2 RAW DATA FROM CGA 72662 LABORATORY STUDIES................. 298
3 HOUSE FLIES KILLED IN FARNAM BAIT FIELD TRIAL.............. 300
4 THE PROBABILITIES, PROBITS, LOG DOSES, AND LOWER AND UPPER
FIDUCIAL LIMITS FROM THE PROBIT ANALYSIS OF SBP-1382
DOSAGE-MORTALITY DATA.................................... 302
5 THE PROBABILITIES, PROBITS, LOG DOSES, AND LOWER AND UPPER
FIDUCIAL LIMITS FROM THE PROBIT ANALYSIS OF BW 21Z DOSAGE-
MORTALITY DATA.......................................... 303
6 THE PROBABILITIES, PROBITS, LOG DOSES, AND LOWER AND UPPER
FIDUCIAL LIMITS FROM THE PROBIT ANALYSIS OF SD 43776
DOSAGE-MORTALITY DATA.................................... 304
7 THE PROBABILITIES, PROBITS, LOG DOSES, AND LOWER AND UPPER
FIDUCIAL LIMITS FROM THE PROBIT ANALYSIS OF ICI ECTIBANTM
DOSAGE-MORTALITY DATA.................................... 305
8 THE PROBABILITIES, PROBITS, LOG DOSES, AND LOWER AND UPPER
FIDUCIAL LIMITS FROM THE PROBIT ANALYSIS OF CARBARYL
DOSAGE-MORTALITY DATA .................... .... ......... 306
9 THE PROBABILITIES, PROBITS, LOG DOSES, AND LOWER AND UPPER
FIDUCIAL LIMITS FROM THE PROBIT ANALYSIS OF MALATHION
DOSAGE-MORTALITY DATA.................................... 307

BiOGRAPHICAL SKETCH.......................... .................... 308












LIST OF TABLES


TABLE PAGE

1. Composition of basal diet for poultry feed trials........... 62

2. Compounds utilized for fly and/or mite control .............. 73

3. Moisture levels(%) of manure samples from the tilling site
and net change in moisture content() ...................... 87

4. Fortified and unfortified diets used during the preliminary
colonization studies with Ophyra aenescens larvae............ 92

5. Combinations of larval and adult diets used during the four
preliminary colonization studies with Ophyra aenescens....... 93

6. Summary of the four Q.:.T'; aenescens adult longevity studies
including average adult life span and the length of the life
cycle from each study....................................... 94

7. The number of pupae, the per cent pupation, the numerical
and per cent emergence, and the larval viability of Ophyra
aenescens reared in fortified and unfortified larval diets.. 97

8. Uneclosed pupal weights of Cr.. : aenescens reared in forti-
fied and unfortified diets.................................. 99

9. Emergence of adults of Ophyra aenescens when various numbers
of first-instar larvae were reared in the same volume of
growth medium.............................................. 100

10. Daily temperatures of larval media as influenced by Cphyra
aenescens larvae over the 7-day larval development period... 102

11. Experimental design of and the larval diet used in the
competition study involving larvae of Ophyra aenescens and
larvae of Misca domestica................................... 103

12. Results of the competition study between Ophyra aenescens
and Risca domestica........................................ 104

13. Experimental design of Ophyra aenescens predation study..... 106

14. Results of Ophyra aenescens predation study.................. 108









15. Experimental designs of and the larval diets used in the
competition studies involving larvae of Hermetia illucens
vs. larvae of Musca domestic and Ophyra aenescens........... 115

16. Experimental design of and the larval diet used in the
competition study involving larvae of Hermetid illucens and
larvae of Sarcophaga robusta................................ 116

17. Results of the competition study between Hermetia illucens
and Ophyra aenescens....................................... 118

18. Results of the competition study between Hermetia illucens
and M usca domestica........................................ 119

19. Results of competition study between Hermetia iZZNcuns and
Sareophaga robusta...................................... .... 122

20. Average daily feed consumption (g/bird per day) of hens fed
diets containing TH 6040 and ZR-515 at 1 and 10 ppm.......... 125

21. Average hen-day production(%) of hens fed diets containing
TH 6040 and ZR-515 at I and 10 ppm.......................... 127

22. Bioassay of manure from hens fed diets containing TH 6040
and ZR-515 at I and 10 ppm.................................. 128

23. Data from pupal traps set in manure from hens fed diets
containing TH 6040 and ZR-515 at 10 ppm..................... 130

24. Number of pupae, number of pupae closed, and per cent
mortality when larvae of Musca domestic were reared in
poultry manure containing two levels of methoprene sand
granules................................ .................... 133

25. Number of pupae, number of pupae closed, and per cent
mortality when larvae of Musca domestic were reared in
poultry manure containing two levels of methoprene sand
granules that were applied at the University of Florida
Poultry Science Farm........................................ 135

26. Fly species and diets used for CGA 72662 laboratory
studies..................................................... 136

27. Summary of per cent larval mortality in CGA 72662
laboratory studies ........................... ............ 138

28. Larval mortality and larviform pupae formation resulting
from various levels of CGA 72662 in growth media of
house flies................................................. 139


PAGE


TABLE









29. The probabilities, log doses, upper and lower fiducial
limits, and probits from the probit analysis of CGA
72662 dosage-mortality data.................................. 142

30. Sectioning of the manure collection area, assignment of
treatments, and the application rates of CGA 72662 and
the organophosphorus larvicides............................. 144

31. Mixing the test concentrations of CGA 72662 and the
organophosphorus larvicides................................. 145

32. Larval population means for all treatments during each
sampling period when poultry manure was treated with
CGA 72662 and three organophosphorus larvicides............. 146

33. Larvicidal activity period of compounds tested in the
CGA 72662 organophosphorus larvicide study.................. 149

34. Weekly treatment means of house fly, soldier fly, and little
house fly populations from manure treated with CGA 72662 and
tilled twice weekly ........................................ 155

35. Treatment and sample collection schedule when CGA 72662 was
added to the drinking water of laying hens as an oral
larvicide................................................... 160

36. Mortality of immature house flies in the manure of laying
hens collected when CGA 72662 was added to the drinking
water at the rates of 10 and 20 ppm.......................... 162

37. Mortality of immature house flies in the manure of laying
hens collected when CGA 72662 was added to the drinking
water at the rates of 1.5 and 5.0 ppm....................... 163

38. Mortality of immature house flies in the manure of laying
hens collected 3 days after treatment of drinking water
with CGA 72662 at 10 and 20 ppm was terminated.............. 164

39. Mortality of immature house flies in the manure of laying
hens collected 5 days after treatment of drinking water
with CGA 72662 at 10 and 20 ppm was terminated.............. 165

40. Monthly catches of Musca domestic, Hermetia illucens,
Stomcxys catcitrans, Hematcbia irritans, and Ophyra sp.
in two blacklight electrocutor grid traps................... 166

41. Results of knockdown tests with Farnam baits................ 170

42. Results of residual tests with Farnam baits................. 171


viii


TABLE


PAGE







TABLE PAGE

43. Treatment means by treatment in Farnam bait field trial..... 173

44. Treatment means by sex in Farnam bait field trial........... 175

45. Results of the knockdown test using BW 21Z and Golden
MalrinTM with MuscamoneTM fly baits..........:............... 178

46. Results of the residual test using BW 21Z and Golden
MalrinTM with MuscamoneTM fly baits.......................... 180

47. Results of the attractiveness test using BW 21Z and Golden
MalrinTM with MuscamoneTM fly baits.......................... 181
TM
48. Results of the field test using BW 21Z and Golden Malrin
with MuscamoneTM fly baits.................................. 182

49. Test concentrations and corresponding responses from the
JFU 5819 laboratory bioassay................................ 184

50. The probabilities, probits, log doses, and upper and lower
fiducial limits from the probit analysis of JFU 5819
dosage-mortality data....................................... 185

51. Mortality and per cent mortality of house flies exposed to
two levels of JFU 5021A applied as a contact residual on
three different surfaces..................................... 188

52. Names, formulations, test concentrations, mixing instruc-
tions, and application rates of compounds applied to
wooden panels............................................... 191

53. Total and per cent mortality that occurred when 3- to 5-day-
old female house flies were exposed to synthetic pyrethroids
on wooden panels............................................ 192

54. Concentrations of acaricides and total, per cent, and
corrected per cent mortality for each concentration tested
against northern fowl mites................................ 195

55. LCso's and regression equations for the acaricides tested... 198

56. The formulations, mixing procedures, test concentrations,
and application rates for acaricides tested at the tilling
site for control of northern fowl mites..................... 201

57. Treatment schedule of acaricides tested at the tilling site
for northern fowl mite control.............................. 202

58. Mite population means and converted population means from
hens treated with malathion, carbaryl, and RavapTM at the
tilling site for northern fowl mite control ................. 203









59. Formulations, mixing procedures, and application rates
for synthetic pyrethroids applied to floor birds in
Chipley, Fl ................................................. 207

60. Pre- and post-treatment field-estimated and converted mite
population counts and treatment means for each treatment
from floor birds treated with two synthetic pyrethroids
in Chipley, Fl.............................................. 208

61. Daily egg production means of birds treated with two
synthetic pyrethroids in Chipley, Fl........................ 213

62. Data collection and treatment application schedule for the
RavapTM northern fowl mite trial in Chipley, Fl............. 217

63. Pre- and post-treatment mite population means by treatment
and strain from caged-layer trial at Chipley, Fl............ 219

64. Pretreatment mite population means by strain (treatment
ignored) and by treatment group from the caged-layer trial
at Chipley, Fl.............................................. 220

65. Post-treatment mite population means by strain (treatment
ignored) and by treatment group from the caged-layer trial
at Chipley, Fl.............................................. 221

66. Means of the combined pre- and post-treatment mite counts
of the control group (treatment 2) from the caged-layer
trial at Chipley, Fl........................................ 222

67. Transformed pretreatment mite population means by strain
(treatment ignored) and by treatment group for the west
end of house 200............................................ 223

68. Transformed pretreatment mite population means by strain
(treatment ignored) and by treatment group for the east
end of house 200............................................ 224

69. Transformed post-treatment mite population means by strain
(treatment ignored) and by treatment group for the west
end of house 200............................................ 226

70. Transformed post-treatment mite population means by strain
(treatment ignored) and by treatment group for the east
end of house 200............................................ 227

71. Egg production means by strain (treatment ignored) and by
treatment group for house 100............................... 228


TABLE


PAGE








72. Egg production means by strain (treatment ignored) and by
treatment group for house 200............................... 229

73. Egg production means by strain (treatment ignored) and by
treatment group for houses 100 and 200 combined............. 230

74. Egg production means by week (treatment ignored) and by
treatment group in the caged-layer trial at Chipley, Fl..... 231

75. T-test of egg production treatment means by strain from
the caged-layer trial at Chipley, Fl........................ 234

76. Egg production means when each quarter of house 200 was
analyzed as a separate treatment............................ 235

77. Egg production means, pre- and post-treatment mite popula-
tion means, and the change in mite population numbers on
untreated hens from the caged-layer trial at Chipley, Fl.... 236

78. Mean operator and tractor time, and the amount of fuel
required to till one 91.4-m California-style poultry
house ...................................................... 245


TABLE


PAGE













LIST OF FIGURES


FIGURE PAGE

1. View of tractor and tiller .................................. 53

2. Tiller in operation........................................ 55

3. Layout and numeric designation of poultry houses at the
tilling site................................................ 56

4. A tagged pupal trap after removal from manure pack.......... 58

5. Light trap opposite egg processing room..................... 65

6. Light trap between egg cooler and house 4................... 66

7. Panel with guttering suspended by chains at the tilling
site........................................................ 68

8. A pair of workers examining a hen for mites................. 70

9. A close-up view of Figure 8................................. 70

10. The appearance of fairly dry manure after tilling........... 77

11. Manure which has dried enough to form particles of various
sizes when tilled........................................... 79

12. Experimental design for adding builder's sand and wood
chips to houses 1 through 4 at the tilling site............. 81

13. The appearance of chips before spreading.................... 82

14. The appearance of chips after the initial tilling........... 82

15. The appearance of manure collection areas at the tilling
site after manure removal and subsequent flooding........... 85

16. Addition of wood chips to flooded manure collection areas... 86

17. Net results of the manure drying experiment with wood chips
and tilling................................................. 88

18. The manure in 3-B at the end of the experiment.............. 90






FIGURE PAGE

19. Graphic representation of the four Ophyra aenescens adult
longevity studies ........................................... 96

20. Regression curve for data from Ophyra aenescens predation
study...................................................... 109

21. Areas on the cephaloskeleton of Ophyra aenescens compared
with those of Musca domestica.............................. 111

22. Basal sclerite of Musca domestica........................... 112

23. Basal sclerite of Ophyra aenescens.......................... 112

24. Oral sclerite of Musca domestica........................... 113

25. Oral sclerite of Ophyra aenescens........................... 113

26. Assignment of diets containing ZR-515 and TH 6040 to
treatment groups in range houses............................ 124

27. Experimental design for testing the effects of ZR-515 sand
granules on larval populations of Musca domestica........... 131

28. A larviform pupa formed in medium containing between 0.5 and
1.0 ppm of CGA 72662........................................ 140

29. Probit curve, fiducial limits, LCso, and regression equation
for CGA 72662 dosage-mortality data.......................... 143

30. Larval population means for all treatments during each
sampling period when poultry manure was treated with
CGA 72662 and three organophosphorus larvicides............ 148

31. Cross-section of manure-wood shavings mixture 1 week after
tilling, showing relative locations of house fly and
soldier fly populations.................................... 152

32. Treatment area, assignment of treatments, and tilling
schedule in the CGA 72662 tilling trial..................... 154

33. Weekly treatment means of house fly populations from manure
treated with CGA 72662 and tilled twice weekly.............. 157

34. Weekly treatment means of soldier fly populations from
manure treated with CGA 72662 and tilled twice weekly........ 158

35. Weekly treatment means of little house fly populations from
manure treated with CGA 72662 and tilled twice weekly........ 159


xiii







FIGURE PAGE

36. Fluctuation in house fly populations as recorded by two
blacklight traps at the tilling site........................ 168

37. Farnam bait field trial treatment means..................... 174

38. Farnam bait field trial treatment means by sex.............. 176

39. Probit curve, fiducial limits, and LCso for JFU 5819
dosage-mortality data....................................... 186

40. Probit curves for all acaricides tested, plotted on one
set of axes................................................. 197

41. Mite population means from hens treated with malathion,
carbaryl, and RavapTM at the tilling site................... 204

42. Pre- and post-treatment field-estimated mite population
means from floor birds treated with two synthetic
pyrethroid compounds at Chipley, F ......................... 211

43. Houses 100 and 200 showing locations of strain replications
and treatment areas......................................... 215

44. Weekly egg production means by treatment from the caged-
layer trial at Chipley, Fl.................................. 232

45. Plot of egg production means vs. precount mite means by
strain from caged-layer trial at Chipley, Fl............... 237


xiv







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


THE EVALUATION OF POULTRY PEST MANAGEMENT
TECHNIQUES IN FLORIDA POULTRY HOUSES

By

Jerome Adkins Hogsette, Jr.

December, 1979

Chairman: Dr. J. F. Butler
Major Department: Entomology and Nematology


The house fly, Musca domestic (L.), and the northern fowl mite,

Ornithonyssus sylviartu (C.& F.), are the two major arthropod pests

associated with the poultry industry in Florida. Presented is a system

whereby various control techniques for these pests have been evaluated.

Techniques were divided into two main areas: house fly control, based

mainly on manure management for control of immature fly populations,

and northern fowl mite control, based on evaluation of acaricides for

control of mite populations on chickens.

Rotovation, a method of tilling and aerating manure for house fly

control, was evaluated as a technique for drying and composting manure

in situ. Drying was enhanced by tilling wood chips and an insect growth

regulator (CGA 72662TM) into wet manure areas. When CGA 72662 was

applied topically, fly larval control was seen for 35 days with a

single application. Commercially labeled organophosphorus larvicides

lasted only 2 weeks. LCs5 bioassay of CGA 72662 for house fly larvae

was 0.45 ppm.

Methoprene and dimilin were evaluated as oral larvicides, but gave

poor field results. When 20 ppm of CGA 72662 were added to the


XV







drinking water of hens, bioassayed manure produced 100% fly mortality.

Methoprene sand granules topically applied to manure gave house ly

larval control of greater than 90% for 3 days post-treatment in bio-

assayed manure samples.

Ophyra aenescens (Wied.), the black dump fly, was reared in the

laboratory and the larvae were proven to be predators of house fly

larvae. In the field, 0. aenescens adults were considered pestiferous,

and their use as a biocontrol agent is not recommended at this time.

In laboratory competition studies, Hermetia illucens (L.), an

assumed biocontrol agent, did not prevent house fly larvae from reaching

maturity when the two species were reared in the same container. In

the field, a situation occurred where larvae of H. illucens and M.

domestic were living in the same manure pack, but in different strata.

Light traps, baits, and residual sprays were evaluated for their

ability to effectively reduce adult house fly populations. Light traps

were generally ineffective, but baits gave good results. In laboratory

studies, a bait consisting of dichlorvos and ronnel had the fastest

knockdown, killing all flies in 10 minutes. A methomyl bait had the

longest residual and was killing at a rate higher than 25% after a
TM
6-week testing period. A Bomyl bait with Lure'em II attractant

killed a significantly greater number of flies than all baits tested.

A permethrin bait was unattractive to flies in the field even though

its fly killing ability was demonstrated in the laboratory. When

ICi 143, BW 21Z, and SD 43775 synthetic pyrethroid compounds were

applied as a residual treatment to wooden panels hung in poultry houses,

ail compounds produced 100' fly mortality 121 days post-treatment.







In laboratory bioassay, permethrin LDso for house fly adults was

18.0 ppm.

Northern fowl mite acaricide bioassays gave LCso's for carbaryl,

malathion, and permethrin of 0.41, 1.70, and 2.9 ppm respectively.
TM
When carbaryl, malathion, and RavapT were applied to hens at rates

of 0.30, 0.44, and 0.36% respectively, northern fowl mite control

approaching 100% was achieved with Ravap in 2 weeks. Adequate control

was achieved after 4 to 6 weeks with reapplication of carbaryl and

malathion. Two synthetic pyrethroids, BW 21Z and SD 43775, applied

once to floor birds at rates of 0.05% and 0.10% respectively, gave

100% control of mites for 7 weeks. In production trials when 12

strains of hens were evaluated for the effects of northern fowl mites

on egg production, no overall difference in egg production could be

found due to mite control. However, one strain of hens showed a

significant increase in egg production of 3.67% due to mite control

with Ravap.

The use of the above techniques, individually or in combinations,

will enable the poultry operator to more efficiently regulate house

fly populations in and around poultry houses. These techniques will

also enable him to effectively control northern fowl mites on hens

and determine whether or not mite populations are affecting hen

performance.


xv i












INTRODUCTION

In many areas of Florida, poultry farms are located in close

proximity to large housing developments. House flies (' usca domestic,

L.) must be effectively controlled for more than just economic reasons,

since poultrymen are quickly blamed when flies are found in or around

nearby homes. Under Florida law, Chapter 386, which regulates ex-

cessive fly breeding and odors, complaints by three responsible citizens

can result in sanitary inspections of suspect poultry farms by Health

and Rehabilitative Services (HRS). Inspected farms not meeting HRS

standards for sanitation and fly control can eventually be closed if

farm owners fail to rectify discrepancies to the satisfaction of HRS and

the surrounding neighbors. Although not a problem for homeowners,

northern fowl mites, Crnithonyssus sylviaruf (C.& F.), car, be annoying

to farm laborers and egg processors. Mites have also been known to

cause decreased egg production and increased mortality in poultry

flocks.

Although many compounds are available for fly and mite control, a

large number have been rendered ineffective due to resistance problems.

Slowness and indecision by the EPA have prevented the labeling of new

compounds and put the labels of approved compounds in jeooardy.

In this dissertation house fly control was evaluated by roto-

tilling, an in situ method for drying poultry manure. Stabilizers were

added to wet manure to enhance drying. Insect growth regulators (IGR's)

were tested in poultry feed and water as orai larvicides. IGR's were

also evaluated when applied topically to poultry manure.

1









Ophyra aenescens (Weid.) was tested as a biocontrol agent against

house flies. Laboratory and anatomical evidence is presented to show

that larvae of 0. aenescens are predaceous. The ability of Hermetia

illucens (L.) to preclude larvae of other flies from its growth media

was investigated and results corroborated by field observations.

Light traps, granular baits and contact residual insecticides were

evaluated for adult house fly control.

The efficacies of labeled and unlabeled acaricides were evaluated

for northern fowl mite control. The effects of northern fowl mites on

egg production were evaluated on 12 strains of laying hens.












LITERATURE REVIEW

The House Fly

History and Economic Importance

The house fly, iilusca domestic (L.), is a major economic pest of

livestock and poultry. In 1977 the poultry industry in Florida lost

an estimated S5.6 million due to flies (Butler, 1979). The mere

presence of house flies in great numbers indicates the need for improved

sanitation measures (Scudder, 1949) and may trigger legal action.

From early Biblical times, when swarms of flies ravaged Egypt,

through ancient history (Cloudsley-Thompson, 1976), and up to the pre-

sent, flies have been noted as pests (Greenberg, 1973). Flies fulfill

all the conditions required of a disease vector (Greenberg, 1971), and

have been rated second only to man as the most important animal in the

transfer of human disease (Scudder, 1949). A single fly may carry more

than 1 million bacterial cells. Any particular fly may be contaminated

with more than 100 species of pathogenic organisms capable of causing

such diseases as dysentery, typhoid fever, cholera, salmonellosis,

anthrax, poliomyelitis, and hepatitis. Flies may also be contaminated

with eggs of nematodes and cestodes (James and Harwood, 1969). Books

by Greenberg (1971, 1973), Hewitt (1914), Lindsay (1956), and West

(1951) should be consulted for in-depth details of fly-borne diseases.

What makes the house fly sc important in these disease transmission

cycles is its close coexistance with man, its consumption of both con-

taminated and uncontaminated food, its great flight activity and

3







dispersal, and its constant alternation between feces and food

(Greenberg, 1971). Besides the transmission of diseases and helminths,

house flies also cause myiasis. Cuticular, ocular, and urinary myiasis

seem to be the types most frequently reported (James, 1947), but other

types are recorded in the literature (Leclercq, 1969b).

Bionomics

Distribution

The house fly (.'(2, domestic) was described by Linnaeus in 1758,

and is known as an ubiquitous insect (West, 1951). Hewitt (1914) called

the house fly a qualified ubiquitous insect because M. domestic is

divided into subspecies groupings in some geographic areas of the

globe. These subspecies are listed in Stone et al. (1965).

Life cycle

The house fly life cycle varies in length depending on the ref-

erence. Bishopp et al. (1915) stated that the entire cycle required

from 7 days to 7 weeks. Other estimates are 3 weeks per generation

(James, 1947), 10 days under usual conditions but 7 days in warm

weather (James and Harwood, 1969), and less than 1 week in the tropics

(Oldroyd, 1965). The following times are given for the length of the

developmental stages by Bishopp et al. (1915) and James and Harwood

(1969) respectively:

Eggs hatch in: 24 hours 10 to 12 hours

Larval stages: 3 days to 3 weeks 5 days

Pupal stage: 3 to 26 days 4 to 5 days

Adult life span: --- 30 to 60 aacs








Some differences in the reported lengths of the life cycle can be

attributed to varience in environmental factors, such as temperature.

Melvin (1934) studied the duration of incubation periods when house fly

eggs were incubated at different temperatures. Incubation periods

ranged from 51.45 hours at 15.0 C to 8.05 hours at 40.6 C. At 42.8 C,

no eggs hatched.

In fresh poultry manure, a temperature of 27.0 C and a moisture

level of 60 to 75% proved optimal for larval development (Miller et al.,

1974). In horse manure, larvae showed no ill effects when the tempera-

ture reached 45.0 C, out as the temperature approached 48.9 C, they

began to migrate out. At 54.4 C, larvae died within I min, and at

60.0 C, death was instantaneous (Allnut, 1926).

The pH of the larval medium may also change cycle length. Erofeeva

(1967) determined the optimum pH for house fly larval media to be between

7 and 8. This is also the pH of day old poultry manure (Beard and

Sands, 1973).

Another temperature dependent variable that changes the length of

the life cycle is the fly's ability to overwinter. Early investigators

were unable to determine in which stage M. domes7ica overwintered

(Hewitt, 1914; Graham-Smith, 1916), but it is now known that the house

fly can overwinter in all of its developmental stages (Greenberg, 1971).

Breeding occurs throughout the year in warmer climates where tempera-

tures are 18.0 C or above (James, 1947; Greenberg, 1971).

Fecundity

At 5 to 7 days of age, the female house fly has mated and is ready

to begin laying eggs (James and Harwood, !969). A female may lay up to








1000 eggs (LaBrecque et al., 1972), and produce 5 to 20 or more batches

of eggs with each batch containing 120 to 150 eggs (James, 1947). Popu-

lation increases of up to six-fold can rapidly occur in field popula-

tions of house flies if conditions are right (LaBrecque et al., 1972).

House flies oviposit all day without regard to time, with no more eggs

laid in the morning than in the afternoon (Meyer et al., 1978).

Larval habitats

Larvae of M. domestic will develop in any decaying and fermenting

organic material (James, 1947), in kitchen refuse and decaying vege-

tables (Bishopp et al., 1915), and in manure of all types (Bishopp et al.,

1915; James, 1947), especially horse manure (Hewett, 1914, James and

Harwood, 1969).

House fly larvae developing in media such as corpses or garbage

cannot be killed by burying the media less than 1.2 m deep. Larvae

will climb to within 30.5 cm of the soil surface, pupate and about 90%

will survive (Mellor, 1919). Larvae that develop in microbial contami-

nated media may produce adults free from contamination (Greenberg,

1973).

As in other families of diptera, males of M. domestic emerge

from their puparia before the females (Mellor, 1919).

Adult dispersion patterns

In most experiments designed to study house fly dispersal patterns,

marked flies were released and recaptured at different intervals from

the release point. Results indicate that flies can disperse from

8.3 (Quarterman et al., 1954) to 20.0 km (Bishopp and Laake, 1921:

Lindquist et al., 1551). Greenberg (1973) reported a dispersion of

2.3 to 11.8 km within 24 hr.







Fiy movement is apparently random (Schoof and Siverly, 1954a),

especially when winds are variable (Pickens et ai., 1967). Flies

tended to disperse upwind when a steady 3.3 to 11.7 kph wind was

blowing and dispersal rate increased when temperatures were 11.7 C or

above (Pickens et al., 1967). A house fly apparently spends most of

its life going from site to site (Schoof and Siverly, 1954a). In the

study by Pickens et al. (1967), flies traveled 0.8 km past a clean

farm to reach a dirty farm.

Nocturnal resting places of adults

Since the advent of contact residual pesticides, detecting and

then treating house fly nocturnal resting places had been advocated as

a method of control (Scudder, 1949). Kilpatrick and Quarterman (1952)

found that flies congregate at dusk in large numbers and stay in the

same place all night. In hot weather they rest outside on vegetation,

but in cooler weather they rest inside structures (Oidroyd, 1965).

Nocturnal resting sites of house flies are usually within 6.1 m of a

favored daytime feeding and breeding area and are usually above the

ground, but rarely higher than 4.6 m (Scudder, 1949). Anderson and

Poorbaugh (1964a) found that on a test poultry farm 85'% of the house

fly population rested inside the poultry houses at night.

Methods for Larval Control in Poultry Houses

Manure management in dry systems

An average size hen produces from 90.9 g (Hart, 1963) to 168.2 g

of wer manure daily (Winter and Funk, 1941). Fresh manure is approxi-

mately 70% moisture (Card and Nesheim, 1975; Hart, 1963) and has a pH

of about 6 that rises to between 7 and 8 after about 12 hr due to









bacterial action (Beard and Sands, 1973). Since house fly larvae pre-

fer a moisture level between 60 and 75% (Miller et al., 1974) and a ph

of between 7 and 8 (Erofeeva, 1967), keeping manure dry and thereby

stabilizing the manure habitat is one of the most important goals in

fly control (Hartman, 1953; Legner et al., 1975; Wilson and Card, 1956).

If moisture levels in manure exceed 80%, manure becomes anaerobic

(Miller et al., 1974), rendering it unsuitable for house fly develop-

ment (Beard and Sands, 1973).

Some authors have advocated frequent manure removal, i.e. every 5

days, to achieve good fly control (Wilson and Card, 1956). Others found

that monthly or bi-weekly manure removal favored fly populations (Peck

and Anderson, 1970). Abstention from manure removal has allowed popu-

lations of predators of dipteran larvae to increase (Peck and Anderson,

1969; Peck, 1969). Axtell (1970) attained good fly control by removing

manure early in the fly season and then using residual sprays to keep

adults in check. Loomis et al. (1975) recommended infrequent manure

removal where drying was enhanced by frequent mechanical stirring. Al-

though mechanical stirring does not succeed in drying manure in all

situations (McKeen and Rooney, 1976), mechanical stirring, or rotova-

tion, has proven to be a successful method for controlling flies on

poultry farms in the Tampa Bay area of Florida (Hinton, 1977). If

manure must be removed from poultry houses, a dry base 12.7 to 15.2 cm

deep should be left to help dry out fresh droppings (Hartman, 1953),

and re-establish house fly predators (Peck and Anderson, 1970).

Other management practices that a poultry farm operator can use to

implement a manure management program are to prevent water from reaching









the manure, increase the drying surface of the manure, improve the

amount and speed of airflow over the manure, and reduce the amount of
2
fresh manure per m of floor space (Hartman, 1953).

One of the benefits of keeping manure dry is that it retains its

value as a fertilizer (Hinton, 1977; Loomis et al., 1975). One of the

drawbacks of using poultry manure as a fertilizer, however, is that

manure can increase soil salinity when it is applied in high levels

(Shortall and Liebhardt, 1975). This is not a problem on acid eastern

soils.

Hammond (1942) may have been the first to formulate diets for

growing chickens using cow manure as a vitamin supplement. Since then,

many types of manures have been tested as feed supplements. Poultry

manure, which has an average nitrogen, phosphorus, and potassium

analysis of 4, 3, and 2%, respectively (Woods, 1975), has been tested

in poultry diets (Lee and Blair, 1973; Lee et al., 1976) and advocated

for use by some authors (Woods, 1975).

For other options in manure management, consult the book on agri-

cultural waste treatment by Hobson and Robertson (1977).

House fly biocontrol agents

The predators and parasites of immature house flies are many.

Since few were dealt with in this study, only a selected review is

deemed necessary. Additional references pertaining to house fly

predators and parasites can be Found in the house fly bibliography of

west and Peters (1973). Books by Askew (1971), Clausen (1940), and

Thompson (1943) are also recommended.








Spalangia endius (Walker)

Due to the individual attention that has been given this pupal

parasite, and since it contaminated several of our fly colonies, a

brief review is warranted.

Spalangia endius, Walker (Hymenoptera: Pteromalidae) was found to

be not only a fairly common pupal parasite of house flies in poultry

manure, but one which could outcompete other species of microhymenop-

terans (Ables and Shepard, 1974; Legner, 1967; Legner and Brydon, 1966).

It is noted for its ability to rapidly find hosts (Ables and Shepard,

1974) and parasitize more hosts per unit time than its competitors

(Legner, 1967; Legner and Brydon, 1966). Best results were achieved

with S. endius during hot, dry weather (Olton and Legner, 1975). Be-

sides the pupae of M. domestic, S. endius also parasitizes pupae of

Fannia femoralis and Ophyra Zeucostoma (Legner and Brydon, 1966).

Morgan et al. (1975) suppressed a population of house flies in 35

days on a north Florida poultry farm using continuous releases of

S. endius, and Weidhaas et al. (1977) designed a model to simulate the

parasite-fly system. Thornberry and Cole (1978) found S. endius to be

effective only on isolated farms with dry manure. Morgan et al. (1976)

performed a laboratory study of the host-parasite relationships of

S. endius and M. domestic and then devised a method for mass rearing

the pupal parasite in the laboratory (Morgan et al., 1978).

Ophyra aenescens (Wied.)

Ophyra aenescens (Wied.) is a shiny black muscid fly easily dis-

tinguished from other members of the genus by its rufous-yellow palpi.

It was described in 1830 by Wiedemann, who placed it in the genus









Anthomyia; in 1897, Stein transferred it to Ophyra Robineau-Desvoidy

1830 (Johnson and Venard, 1957). Several subsequent descriptions of

the genus have been published, all placing Ophvyr, in the family

Anthomyiidae (Malloch, 1923; Seguy, 1923; Aldrich, 4928; Graham-Smith,

1916; and Bryan, 1934). After studying the male terminalia, Crampton

(1944) decided that Ophyra was a typical muscid, and placed it in that

family, where it remains. Sabrosky, in 1949, described the genus in

the Pacific region.

Distribution. tOhyra aenescens occurs in the United States from

Oregon to Arizona, and from Illinois to the East Coast and Florida

(Greenberg, 1971). It is also found in the Neotropics, the Galapagos

Islands, Hawaii, Nauru, the Ocean Islands, and possibly in Bermuda

(Stone et al., 1965).

Biology and rearing. The biology and morphology of Ophyra

aenescens were described by Johnson and Venard in 1957. They used a

larval medium consisting mainly of C.S.M.A.TM standard preparation.

Initially, adults were maintained on cane sugar dissolved in water, but

no fertile eggs were produced until a source of animal protein was pro-

vided. Fish meal was used dry as a protein source and moistened as

a site for oviposition. Eggs hatched in 12 to 16 hours at 280 C.

The development periods for the three larval instars and the pupal

stage averaged 9 and 4 days, respectively. The complete cycle required

a minimum of 14 days at 270 C + I. Males lived an average of 15 days

and females lived an average of 20 days.

Roddy (1955) used a bacto-agar medium for rearing larvae. Prepara-

tion was time consuming and laborious compared to the C.S.M.A. medium of

Johnson and Venard (1957).







Predaceous nature. Hobby (1934) suggested that adults of Ophyra

might be predaceous. He noted them apparently feeding on dead insects,

but did not see them actually capture prey.

Seguy (1923) stated that the larvae of Opihyra are predaceous. This

was supported by Keilin and Tate (1930) who described the larvae of

0. leucostoma as having buccopharyngeal armature characteristic of

larvae that are both saprophagous and carnivorous. Later experiments

proved that 0. leucostoma was predaceous (Peck and Anderson, 1969;

Peck, 1969), but not cannibalistic (Anderson and Poorbaugh, 1964b).

Relationship with Musca domestic in poultry manure. Ophyra

Zeucostoma (Wied.) occurs commonly in poultry houses in many parts of

the world (Peck and Anderson, 1970; Legner and Olton, 1968; Fujito

et al., 1966). Ophyra capensis (Wied.) has been reported from poultry

houses in Britain (Conway, 1970 and 1973), and 0. aenescens from

poultry houses in Florida (P. G. Koehler, personal communication).

Hermetia illucens (L.)

Description. Hernmetia illucens, the black soldier fly (Sutherland,

1978), is a rather large hemisynanthiopic stratiomyid fly that is easily

recognized (Greenberg, 1971). The genus Hermetia can be distinguished

from all other North American genera of Stratiomyidae by the length of

the style of the flagellum, which is as long as or longer than the

remaining segments of the flagellum (James, 1935).

Linneaus described H. iZlucens in 1738 (May, 1961). Malloch (1917),

Ricardo (1929), and Borgmeier (1930) described the immatures and pupae.

James (1935), Linder (1938), and lide and Mileti (1976) described the

adults, with Linder's description being the most detailed. May (1961)

described both adults and immatures.









Bionomics of Hermetia. The eggs of H. illucens take between 5 and

14 days to hatch at room temperature (May, 1961). They are laid singly

to form masses of 500 to 1000 eggs (Furman et al., 1959). As many as

1062 may be laid by one female (May, 1961).

Larvae have been reared by placing the eggs in either moistened

C.S.M.A. standard larval fly medium (Furman et al., 1959; Tingle et al.,

1975), or in a medium consisting of dried milk, yeast, water, and paper

tissue (May, 1961).

Larval development at 27 to 28 C required a minimum of 31 days

(May, 1961). There are six larval instars as determined by measurement

of molted head capsules. The first four instars have a creamy appear-

ance, but a day or two after molting occurs, the cuticle of the fifth-

instar larvae becomes shagreened and darkens to greyish yellow. The

cuticle darkens even more after the molt to the sixth instar.

Before pupation, the larvae arrange themselves in a vertical manner

in the medium with the head protruding above the surface and the two

posterior segments curved ventrally (May, 1961). Furman et al. (1959)

reported a pupation period of about 2 weeks at 21 to 28 C, but several

pupae closed after 2 to 5 months. The cycle from egg to adult required

38 days at about 29.3 C in greenhouse conditions (Tingle et al., 1975).

Furman et al. (1959) demonstrated that the larvae of H. illucens

are not paedogenic. Larvae fed on dead larvae and adults, but were

not predaceous or cannibalistic.

Both Furman et al. (1959) and Tingle et al. (1975) found adults of

H. il~'2cns to be eurygamous. The adults reared by the former authors

did not mate, but the females laid masses of sterile eggs. Tingle et ai.








(1975) succeeded in getting H. iZZucens to mate by placing adults in

large (76 x 114 x 137 cm) cages directly in the sun. Few matings

occurred during cloudy weather or when the insects were shaded. Mating

commenced during flight as stated by Copeilo (1926).

Due to the variable length of the larval and pupal stages, there

are probably no more than two generations of H. illucens produced in a

year (Copello, 1926), with overwintering occurring in the larval stage

(May, 1961). Greenberg (1971) states that the adults readily enter

houses while Furman et al. (1959) claim they do not.

Distribution. Hermetia itZucens is rather widely distributed

throughout the Western hemisphere, the Australian region from Samoa to

Hawaii, and in some areas of the Palearctic region (Greenberg, 1971).

Various authors report the presence of H. itlucens in the Eastern

hemisphere (Barbier, 1952; Peris, 1962; Adisoemarto, 1975). James

(1935) states that H. iZZucens has been spread by commerce. Van Dyke

(1939) believes H. iltucens is a European species, but Leclercq (1966,

1969a) claims it is an American species transported to Europe and Asia.

Larval habitats. Immature stages of H. illucens are found in a

variety of habitats. Copello (1926) found them living in beehives in

Argentina where the larvae were destroying the weaker hives. Van Dyke

(1939) found larvae of H. illucens in honey bees' nests in the U.S.

Larvae have also been reported from nexts of Melponidae, a family of

stingless bees (Borgmeier, 1930), from dead crabs (Ricardo, 1929), and

from a human cadaver (Dunn, 1916). Other habitats include beeswax,

catsup, decaying vegetables, potatoes (Malloch, 1917), and outdoor

privies in the Southern U.S. (James, 1947).








Myiasis. Larvae of H. iZZucens may cause myiasis in man, particu-

larly intestinal myiasis due to accidental ingestion of eggs or larvae

(James, 1947; Greene, 1952; Werner, 1956).

Predators and parasites. Only one predator of'H. illucens is noted

in the literature. Bodkin (1917) found specimens of H. illucens in the

nests of Bembecid wasps in British Guiana.

Wasps in the family Diapriidae are the only ones known to parasi-

tize pupae of H. illucens. One species of Diapriid was found by Costa

Lima and Guitton in 1962, and another, Trichopria n. sp., by Mitchell

et al. in 1974. The latter parasite was reared (Tingle et al., 1975)

and had an average life cycle of 26 days at 26.8 C. An average of 86

parasites emerged from each parasitized pupa. Twenty-three per cent

of the field-collected pupae of H. illucens were parasitized (Tingle

et al., 1975).

Relationship with Musca domestic (L.) in privies. The presence

of larvae of H. -ilucens and M. domestic in privies is well documented

in the literature (Howard, 1900; Hewitt, 1914; Parker, 1918; James,

1947; Quarterman et al., 1949; Schoof and Siverly, 1954b; Kilpatrick

and Bogue, 1956). Further studies of the fly-breeding conditions in

privies revealed an apparent antagonistic relationship between the

larvae of these two species. When extremely high numbers of H. illucens

larvae were found in privies, no larvae of M. domestics were present

(Fletcher et al., 1956). Hypothesizing that the larvae of H. iliicens

may interfere with the development of M. domestic, a laboratory test

was performed where various numbers of larvae of both species were

grown together and separately in C.S.M.A. standard larval media.








Musca domestic adults emerged in approximately the same numbers from

all jars and it was concluded that no antagonistic relationship existed

(Fletcher et al., 1956).

Kilpatrick and Schoof (1959) noted thar larvae pf I!. domestic were

absent from privies where excretia was semiliquid and infestations of

H. illucens were heavy. Attempts to dry the excretia with sawdust or by

water manipulation caused excretia to crust over and resulted in an in-

crease of house fly breeding and a decrease in soldier fly breeding.

Relationship with Musca domestic (L.) in poultry manure. The

presence of larvae of M. domestic and H. illucens in poultry manure is

also well documented in the literature (Cunningham et al., 1955; Tingle

et al., 1975). The latter authors found them in Florida and claimed

that the house fly population at one farm was being controlled by the

soldier fly population. Few details were given to support that claim.

The hypothesis that larvae of H. illucens and M. domestic are

antagonistic was again tested in the lab for Furman et al. (1959). This

time, larval house fly populations did not develop in culture medium

containing soldier fly larvae. Neither this experiment nor the previous

one (Fletcher et al., 1956) had treatment repetitions and discrepancies

do exist.

In the field, it was shown that H. illucens larvae will replace

M. domestic larvae in poultry manure if the manure is moistened (Furman

et al., 1959). It was also demonstrated that larval populations of H.

i.lucens will develop successfully when the larvae are introduced under-

neath the crust of dry manure.








The outlook for H. illucens as a biological control agent in Mexico

is considered good (Vazquez-Gonzalez et al., 1962). These authors advo-

cate keeping poultry manure wet, especially in the dry season, and

destroying manure cones to augment H. il'ucans populations.

Chemical control. In the past, most of the chemicals used for fly

control in privies were shown to cause resurgence of house fly popula-

tions and damage soldier fly populations (Kilpatrick and Schoof, 1959).

Under normal circumstances, privies produced few house flies. This was

attributed to water content of the excretia and the presence of H.

illucens. When privies were sprayed with dieldrin, BHC, or chlordane,

house fly production greatly increased. DDT, malathion, and diazinon

had little or no effect on house fly production.

Axtell and Edwards (1970) field-tested various larvicides against

larval populations of H. illZucens in poultry manure. The best control
TM
was achieved with a 0.5% solution of Ravap After eliminating the

soldier fly populations, retreatments were necessary to control resur-

gent house fly populations.

House fly pathogens

Bacillus thurengiensis has been fed to caged layers for fly control,

but when fed at levels providing the best control, decreases in feed

consumption, body weight, and egg production resulted (Burns et al.,

1961). When sprayed on manure as a larvicide, B. t'hirengiensis was

effective against fly larvae and did not damage populations of preda-

ceous mites (Wicht, Jr. and Rodriguez, 1970). Records of other types

of pathogens affecting house flies are abundant in the literature

(Briggs and Milligan, 1977; Burges and Hussey, 1971; Kramer, 1964; Beard

and Walton, 1965).








Insect growth regulators (IGR's)

The first juvenile hormone was extracted from the abdomen of a

male cecropia moth over 20 years ago (Williams, 1956). Researchers have

since been trying to develop compounds showing juvenile hormone activity

for use as pesticides that would be specific for limited species of

target insects but would not be detrimental to the environment (Novak,

1975). House flies were sensitive to the early IGR's (Herzog and Monroe,

1972) as were mosquitoes (Spielman and Williams, 1966). Several books

are available giving the history, chemistry and mode of action of IGR's

(Novak, 1975; Gilbert, 1976; Menn and Beroza, 1972), and the types of

compounds exhibiting juvenile activity on insects (Slama, 1971).

Methoprene

Methoprene, or ZR-515, has been widely tested for the control of

mosquitoes, house flies, and other diptera. Treatment residuals are

rapidly degraded by sunlight and the half life is only 2 to 24 hours

depending on the type of formulation (Schaefer and Dupras, 1973).

Methoprene does not leach out of treated media into the environment

(Wright and Jones, 1976) and is not active against nontarget insects

in bovine fecal pats (Pickens and Miller, 1975).

As a feed additive, methoprene gave significant fly control when

fed to cows at 2.5 mg/kg (Miller and Uebel, 1967). Breeden et al. (1968)

fed methoprene to chickens in 86.9% technical and 7% encapsulated formu-

lations. The technical formulation at 50 and 100 ppm gave good fly

control 3 days and I day post-treatment respectively. The encapsulated

formulation at 5 and 10 ppm gave good control 8 and 2 days post-treatment

respectively. Adams et al. (1976) fed methoprene to hens for 42 days at








10 g/ton of feed. Good larval control was achieved, but inability to

produce total control was blamed on migration of adult flies. Morgan

et al. (1975) found that methoprene in chicken feed at 0.0005 and 0.01%

produced mortalities of 70.9 and 99.3%, respectively, and had no effects

on the hens' weight. Methoprene was not effective, however, when

poultry manure was treated topically in the field.

Dimilin

Dimilin, also known as TH-6040 and diflubenzuron, has been classi-

fied as an inhibitor of chitin synthesis. Many analogues of dimilin

have been synthesized and tested, but none are as effective as difluben-

zuron itself (DeMilo et al., 1978). In the larval stages, dimilin

causes rupture of larval cuticle during or shortly before the next molt

(Jacob, 1973). Topical application to pupae can affect emergence of

adults (Cerf and Georghiou, 1974). Application of dimilin to house fly

adults can result in the suppressed hatchability of eggs laid long after

the application date (Wright and Spates, 1976).

Even though dimilin was active against all major nontarget insects

in bovine fecal pats (Pickens and Miller, 1975), poultry farms treated

topically with dimilin had greater parasitoid populations and species

variety than did farms treated with dimethoate (Ables et al., 1975).

When dimilin was fed to chickens at 6.2 to 12.5 ppm, fly control of

100% was achieved, but residues were found in all eggs sampled (Miller

et al., 1975).

Resistance to IGR's

House fly resistance has been demonstrated for both methoprene and

dimilin (Plapp and Vinson, 1973; Oppenoorth and Van Der Pas, 1977;

Georghiou et al., 1978).







Chemical larvicides

The use of chemical pesticides started about the same time the

poultry industry began keeping chickens in cages (Hartman, 1953). The

following is a brief review of chemicals that have been used as larvi-

cides in poultry manure and their efficacy at the time they were tested.

For a more complete review of larvicides, see Miller (1970).

The idea of oral larvicides evolved in the late 1920's. Cows were

fed tannic acid, linseed oil, Mg2SO,,, and NaCI as possible controls for

horn flies (Miller, 1970).

Wolfenbarger and Hoffmann (1944) may have been the first to advocate

the use of DDT as a house fly larvicide on poultry farms. An emulsion

of 0.25% DDT applied to manure at 1.9 1/9.3 m2 gave good house fly

control, but soldier flies, Hermetia illucens, were fairly tolerant

(Tanada et al., 1950).

A 1% solution of malathion EC applied at 3.8 1/9.3 m2 controlled

fly larvae after two applications 5 days apart. Adults resting on

manure were also killed (Mayeux, 1954a). Malathion was more toxic to

predatory mites than to house fly larvae (Axtell, 1966).

Diazinon applied as a liquid and as a dust controlled fly larvae

for 1.5 to 2 weeks, but fly resurgence occurred after 2 weeks (Wilson

and Gahan, 1957). Wicht, Jr. and Rodruguez (1970) achieved good control

with diazinon and claimed little damage was done to predatory mite

populations. Axtell (1966), however, reported that diazinon is just as

toxic to mites as it is to flies.
TM
Dichlorvos, 20% Shell VaponaT resin strips ground up, gave good

control of house fly larvae and adults for about 7 weeks with three








treatments (Bailey et al., 19716). Dichlorvos is also toxic to predaceous

mites (Axtell, 1966).
TM
Rabon when applied to poultry manure as a larvicide, controlled

flies for 1 (Bailey et al., 1968) to 2 weeks (Matthyssee and McClain,

1973). Rabon was also fed to dairy cows as an oral larvicide (Miller

et al., 1970), but it proved to be ineffective in commercial operations

(Miller and Pickens, 1975).

Thiocarbamide or thiourea, when applied weekly to manure at a rate

of 0.26 g per bird in 152.0 1 of water, achieved between 68 and 94%

control of fly larvae (Jaynes and Vandepopuliere, 1978). Thiourea, as

a larvicide, affects first-instar larvae more than second-instar larvae,

and second-instar larvae more than third-instar larvae. Fly eggs and

pupae are not affected (Hall et al., 1979).

Methods for Adult House Fly Control in Poultry Houses

Light traps

Ultraviolet light between 3300 and 3700 angstroms is effective for

attracting flies (Tarry et al., 1971). Claims of good control of flies

with light traps, however, are sometimes the results of tests performed

with small fly populations (Tarry, 1968), or in ideal situations (Tarry

et al., 1971). Schreck et al. (1975) limited light trap catches to

Stomoxys calcitrans by using CO2 as an additional attractant. Traps

tested by Morgan et al. (1970) averaged 439.1 house flies per day over a

22-day period. Pickens et al. (1975) increased the house fly catch

2.4 times by placing a heated fly bait in the trap.

Trap height influences fly catches. Pickens et al. (1975) found

that lowering traps from ceiling level to 0.5 m above the ground








increased the house fly catch 1.8- to 4.6-fold. Driggers (1971) caught

10.24 times as many house flies with traps at ground level than with

traps 1.5 m above the ground. Prime trapping time for house flies at a

north Florida poultry farm was from 5 min before sunset to 5 min after

sunset (Driggers, 1971). Driggers (1971) reduced house flies at the

farm by 52.8 and 73.1% in 1 and 4 weeks respectively, by using four

light traps placed at ground level in a 121.9 m poultry house. Thimijan

et al. (1972) estimated that 52 light traps would be needed in a

screened dairy barn to capture 0.5% of the 2500 to 5000 flies that were

being released in the barn daily during the test period.

Catches of flies by light traps have been found to be highly

variable. As a result, light traps are recommended for survey work, but

they are not considered consistent enough to accurately estimate fly

populations (Pickens et al., 1972). A more complete summary of light

trap evaluations has been prepared by Hienton (1974).

Baits

An early account of killing flies by attracting them with baits

was published by Morrill (1914). He gives a full account of all items

tested and their efficacy. The best combination was overripe banana

on sticky fly paper.

Most baits used today are granulated sugar baits with or without

attractants. Baits in other forms have been tested with some success.

Mayeux (1954a) made a 1% solution of malathion in honey. Burlap was

painted with this solution and hung in poultry houses to kill flies.

Good control was attained and the bait was active for 1 to 24 days.








Wicht, Jr. and Rodriguez (1970) mixed LC95 concentrations of naled

and ronnel with one-to-one mixtures of malt and water. These solutions

were painted onto squares of waxed paper which were attached to bait

stations made of plywood squares. Paper was replaced weekly. The

naled bait attracted more flies and had a quicker knockdown than ronnel.

Granular baits are convenient to store and use, and have been

tested more extensively than other types of baits. Mayeux (1954b)

reduced house fly populations by 90% or more within 1 hour with a 1%
2
malathion bait applied at 85.2 to 113.6 g/9.3 m If kept dry, the bait

killed at this level for 3 to 7 days. Sampson (1956) ranked the

efficacy of granular test baits in the following order: endrin, hepta-

chlor, lindane, and parathion (all at 0.125%) more effective than

diazinon, dieldrin, DDT, and phenthiazine (all at 0.125%) more effective

than aldrin and thiourea (both at 1.0%). Bailey et al. (1970) tested

1% sugar baits of dimethoate, fenthion, formothion, naled, ronnel, and

trichlorfon. All gave better than 75% control for 18 days.

In 1971, resistance to trichlorfon (from 2.5 to 135.0 times) and

dichlorvos baits (from 2.3 to 16.6 times) was reported from Florida

(Bailey et al., 1971d).

Rogoff et al. (1964) demonstrated the presence of a house fly sex

pheromone which Carlson et al. (1971) later identified as (Z)-9-

tricosene, or MuscalureTM. Muscalure and its homologs were then syn-

thesized in the laboratory by Richter and Mangold (1973). The addition

of Muscalure to sugar baits increased house fly catches (Carlson and

Beroza, 1973). Only males were caught in laboratory studies, but equal

numbers of males and females were caught in the field. Mulla et al.








(1977) tested compounds attractive to house flies and found that

trimethylamine and indole were the main house fly attractants. Baits

consisting of trimethylamine, indole, NH4C1, and linoleic acid were

significantly superior to commercial preparations containing (Z)-9-

tricosene.

Location of bait stations in and around poultry houses was found

to influence the size and sex ratio of the catches. Baits located in

the sunlight-shade border areas collected the greatest number of flies

(Willson and Mulla, 1973). In bait stations near the center aisles,

females outnumbered males, but a one-to-one ratio was approached in

catches from the perimeters of poultry houses (Willson and Mulla, 1975).

Bait stations dominated by one sex had catches significantly lower than

those of stations conducive to both sexes.

Space sprays

This brief review is limited to use of synthetic pyrethroids as

space sprays. In a study by Willis and Thomas (1975), pyrethroids gave

better results than the ronnel standard, and resmethrin gave better

results than allethrin. In another study, ronnel was more effective

than resmethrin (Wilson et al., 1975). Other trials have shown that

resmethrin is effective as a space spray against house flies (Mathis

et al., 1972) and mosquitoes (Haskins et al., 1974). Permethrin was

shown to have a knockdown 8 to 16 times faster than that of allethrin,

and an LD50 three and four times higher than those of mesrethrin and

synergized mesrethrin respectively (Lhoste and Rauch, 1976).

Kissam and Query (1976) tested an automatic piped-aerosol system

that used a 0.71 synergized pyrethrin solution for fly control in








poultry houses. The system provided effective fly control and cost in

the range of other fly control systems.

Contact residuals and resistance

The ability of insects to develop resistance was questioned by

Melander (1914). The question was answered when DDT resistance was re-

ported from several countries in Western Europe in 1947 (West, 1951).

One year later, DDT resistance was reported in the U.S. (Hansens et al.,

1948). DDT had only recently been advocated for use on poultry farms

despite its slow knockdown and kill (Wolfenbarger and Hoffmann,1944).

A survey in Canada showed that house flies were still highly resistant

to DDT (Batth and Stalker, 1970).

Sequential formation of resistance to contact residuals

In 1953, Hansens reported that lindane, methoxychior, chlordane,

and dieldrin applied as residual sprays failed to give control of house

flies. The residual action of diazinon extended 10 weeks against sus-

ceptible flies and 4 weeks against resistant flies (Hansens and Bartley,

1953). Resistance to diazinon was noted soon afterward (Hansens, 1958)

and in Florida it was reported to be 5- to 38-fold (LaBrecque et al.,

1958). By 1970, diazinon resistance was 8- to 62-foid in New Jersey

and a 1% solution failed to give satisfactory control (Hansens and

Anderson, 1970). Flies showing resistance to diazinon also showed re-

sistance to stirofos (Pickens et al., 1972), DDT, methoxychlor, chlor-

dane, dieldrin. lindane, parathion, malathion, dicapthon, ronnel,

trichlorfon, and conmaphos (Hansens, 1958).

Malathion resistance in Florida was about 4-fold in 1956 (LaBrecque

and Wilson, 1961), 133-fold in 1958 (LaBrecque et al., 1958), and 275-

fold in 1960 (LaBrecque and Wilson, 1961).








Hansens and Anderson (1970) found that a 1% solution of the follow-

ing insecticides failed to give satisfactory fly control when applied

as contact residuals: dimethoate, ronnel, stirofos, and bromophos.
TM
FicamTM gave good results as a contact residual against house flies.

Sucrose was added to the solution to improve the knockdown. No resis-

tance data areavailable (Lemon and Bromilow, 1977).

Synthetic pyrethroids

The first synthetic pyrethroid to be synthesized was allethrin

(Schechter et al., 1949) followed by resmethrin (Elliot et al., 1965).

Although natural pyrethrins are known for their quick knockdown (O'Brien,

1967), resmethrin proved to be 55 times more toxic to adult females of

M. domestic than mixed esters of natural pyrethrins (Elliot et al.,

1967). Haskins et al. (1974) claim resmethrin to be effective as a

contact residual, but Mathis et al. (1972) claim the opposite. Syner-

gised resmethrin had increased toxicity against resistant flies and the

synergist prevented knockdown recovery (Schulze and Hansens, 1968).

Decamethrin is a highly toxic pyrethroid ester with an acute oral

LD50 for female rats of 31 mg/kg. It can be rapidly absorbed by in-

halation (Kavlock et al., 1979).

Permethrin is more effective at lower temperatures (Harris and

Kinoshita, 1977). Half life of permethrin in soils with low and high

organic content was 7 and 16 weeks respectively, with the loss of in-

secticide being attributed to microbial action (Williams and Brown,

1979). As in insects, the cis-permethrin isomer was more toxic to

aquatic arthropods than the trans-isomer (Zitko et al., 1979).








The mode of action of pyrethroid poisoning is fairly complex.

Initial signs in insects are usually incoordination and locomotor

instability which are collectively termed knockdown. Details can be

found in Wouters and van den Bercken (1978).

Resistance to pyrethroids can be detected in house flies after

several months of strong selection pressure (Keiding, 1976). Permethrin

resistance has been reported in culicids (Priester and Georghiou, 1978),

and cross-resistance has been reported in DDT-resistant strains of

culicids (Prasittisuk and Busvine, 1977) and cattle ticks, Boophilus

microplus (Nolan et al., 1977).

Shono et al. (1978) reported that metabolic detoxification by

ester hydrolysis and hydroxylation is a major factor limiting the

insecticidal activity of the permethrin isomers.

Northern Fowl Mites

Description and Biology

Economic importance

The northern fowl mite is considered to be the most serious ecto-

parasite of poultry in the state of Florida (L. W. Kalch, personal

communicationn, as well as the U.S. (Sulzberger and Kaminstein, 1936;

Miller and Price, 1977; Smith, 1978). Since it was first recognized as

a poultry pest by Wood in 1920, the northern fowl mite continued to

spread across the country with increasing incidence (Linkfield and

Ried, 1953).

Lyon (1975) stated that in 1970, the northern fowl mite could be

costing the poultry industry $80 million annually. Smith (1978) quoted

DeVaney as estimating an annual $66 million loss due to external








parasites causing decreases in egg production; parasite prevention might

cost as much as $1.1 million. In Florida, Butler (1979) attributed a

$3.7 million loss in poultry profits to the northern fowl mite in 1978.

Taxonomy

Although fowl mites were reported in the literature as early as

1824 (Toomey, 1921), the first accepted name, Dermanyssus sylviarum

(Canestrini and Fanzago), was not seen until 1877 (Cameron, 1938). The

inability of authors to properly identify the northern fowl mite resulted

in the appearance of many synonyms. Several authors have followed this

synonymy through the years until the accepted scientific name of the

northern fowl mite was changed to Ornithonyssus sylviarurn (C.& F.) in

1963 (Cameron, 1938; Furman, 1948; Furman and Radovsky, 1963; Laffoon,

1963).

The northern fowl mite was originally placed in the family

Dermanyssidae, but was later separated to the Macronyssidae (James and

Harwood, 1969). For years, 0. sylviarum was confused with another

poultry pest, the chicken mite, Dermanyssus gallinae. The two can be

distinguished by the shapes of the anal plates and by the shapes of the

dorsal shields (Lapage, 1956; Baker et al., 1956; Weisbroth, 1960).

Ornithonyssus sylviarum has a teardrop-shaped anal plate and the dorsal

shield tapers posteriorly; D. gallinae has a truncate anal plate and the

dorsal shield is more rounded posteriorly. The complete morphology of

the northern fowl mite is well documented (Allred, 1970; Georgi, 1974;

Pound and Oliver, 1976; Krantz, 1978).

Bionomics

Wood (1920) and Cleveland (1923) published early works describing

the biology and life cycle of the northern fowl mite. Cameron's research








(1938) was fairly complete at the time, but since he could not colonize

the mite past the larval stage, he could not fully describe the life

cycle. Colonization has since been accomplished (Chamberlain and Sikes,

1950; Cross, 1954; Cross and Wharton, 1964), and the-entire life cycle

has been described (Sikes and Chamberlain, 1954; Soulsby, 1968). Accord-

ing to Sikes and Chamberlain (1954), females lay an average of two to

three eggs, each one within 48 hours after a blood meal. Eggs hatch in

less than 1 day to six-legged non-feeding larvae which molt in less than

I day to eight-legged protonymphs. Protonymphs take an average of 2

days in which to require the two blood meals necessary for full engorge-

ment. Protonymphs molt to non-feeding deutonymphs that molt to adults

in about a day and a half. Time from adult engorgement to second genera-

tion adult was about 5 to 7 days at 38 to 40 C with a relative humidity

of 90 to 100'. Length of the cycle varies at least partly due to the

intermittent feeding habits of the mites (Cameron, 1938).

The whoie life cycle of 0. sylviarum occurs on the host (Sikes and

Chamberlain, 1954; Kirkwood, 1968; Loomis, 1978); however, oviposition

may occur in the nest of the host (Cameron, 1938). Even though the

northern fowl mite has long been considered a winter pest (Loomis, i?78)

mites have been found on chickens all year round (Kirkwood, 1963 and

1968), and will come out to the tips of the feathers in hot weather

(Cameron, 1938). When separated from the host, 0. s2 vlar-i will Ilve

from 2 to 4 weeks (Cameron, 1938; Baker et al., 1956; Kirkwood, 1963:

Loomis, 1978), as compared to 34 weeks for mc ,rr .:' ? I. e

(Kirkwood, 1963).








The area on the host most preferred by the northern fowl mite is

the vent region (Cameron, 1938), but in severe infestations, mites can

be found over the entire body (Anonymous, 1959; Metcalf et al., 1962;

Loomis et al., 1970). Cameron (1938) seldom found mites on young birds.

Kirkwood (1968) also found this to be true and suggested that it may be

due to lack of contour feathers. He and others (Cameron, 1938; Abasa,

1965) stated that roosters have more mites than hens, possibly due to

differences in plummage. Males have more contour feathers near the vent,

while females have more down near the vent. Feathers are preferred

over down by 0. sylviarum (Kirkwood, 1968).

Cameron (1938) described the erratic behavior of mite populations

on poultry. Mites transfer from bird to bird and populations rapidly

rise and decline, but some birds remain entirely free of mites. This

phenomenon has been seen by other authors (Kirkwood, 1963; Loomis et al.,

1970), who were also unable to explain its cause. Hall and Gross (1975)

found that roosters with high levels of plasma corticosterone response

to social stress that were maintained at high levels of social stress

had lower mite populations than when the conditions were reversed.

Inherited levels of corticosterone had more effect on mites than did

stress alone. It was also found that hens subjected to higher social

stress had significantly lower mite populations than unstressed hens

(Hall et al., 1978; Turner, 1978). Additional experiments indicated

that although hens first coming into production are most susceptible to

northern fowl mite infestation, estrogen alone is probably not responsi-

ble for the difference in mite susceptibility between hens and roosters

(Hall et al., 1978).








Distribution

What were probably the first and the earliest samples of the

northern fowl mite in the U.S. were described by Banks (1906) from

specimens collected in 1895 in North Carolina. Since then, the northern

fowl mite has been found in most of the warmer areas of the U.S. and

Mexico (Benbrook, 1965; James and Harwood, 1969). Some claim northern

fowl mites are found world-wide in the plummage of chickens (Baker et

al., 1956). Citings from Great Britain (Taylor, 1930), Hawaii (Garrett

and Haramoto, 1967), and New Zealand (Thomas and Watson, 1958) sub-

stantiate this claim.

Hosts and methods of dissemination

The northern fowl mite occurs on at least 22 species of birds and

domestic poultry (Benbrook, 1965), and Avian hosts are considered to be

the true hosts (Cameron, 1938). Many papers cite records of northern

fowl mites found on species of native wild birds (Boyd et al., 1956;

Hanson et ai., 1957; Foulk and Matthysse, 1965; Phillis and Cromroy,

1972; Phillis et al., 1976) ana exotic caged birds (Sulzberger and

Kaminstein, 1936; Anonymous, 1951). Several host lists are available

(Peters, 1933; Cameron, 1938; Strandt.mann and Wharton, 1958).

Cameron (i938) lists rodents and man as accidental hosts. Other

such hosts are rabbits (Sikes and Chamberlain, 1954), the house mouse,

Mus muscuZus (Orummond, 957), the big brown bat, Eptesicus ftscl's, the

cave bat, Votis e'siffer (George and Strandrmann, 1960), and the norway

rat, .attus norweg-ius (Hall and Turner, 1976; Miller and Price, 1977).

The northern fowl mite could not be induced to feed on man in the lab

(Sikes and Chanrt' rlain, 1954).








Dissemination studies are few. Besides spreading from bird to bird

(Cameron, 1938), Foulk (1964) found that four main methods of poultry

flock infestation are by infested hatcheries and contract started-pullet

farms, infested trucks and crates used :o carry infested birds, infested

personnel, equipment, or egg crates, and infested wild birds that enter

poultry houses. While Hartman (1953) believed northern fowl mites to

be carried by sparrows, Foulk (1964) was unable to infest chicks with

northern fowl mites from sparrows. Mites have also been carried from

farm to farm on filler flats that have not been fumigated after use

(Anonymous, 1968).

Since the northern fowl mite has been found on the Norway rat and

the house mouse in poultry houses (Hall and Turner, 1976; Miller and

Price, 1977), it is assumed that these rodents may aid in mite dis-

semination.

Effects on the Host

Patent effects of northern fowl mite infestation

The most obvious sign of a northern fowl mite infestation is

feathers in the vent area which have become matted and discolored

(Yunker, 1973) from the eggs ana excretion of the mites (Metcalf et al.,

1962). Examination of birds reveals mites and usually evidence of

skin irritation and feather plucking (Anonymous, 1967). In more severe

cases, the skin becomes thickened and scabby (Anonymous, 1959; Metcalf

et al., 1962; Yunker, 1973) due to secondary infection of the bites

(Cameron, 1938).

While northern fowl mites seen crawling on freshly laid chicken

eggs are an indication of a mire infestation, the number of mires








observed is not necessarily an indication of the severity of the infes-

tation (J. F. Butler, personal communication). The way to determine

the severity of infestation is to directly examine the suspected fowl

and check for the symptomology described above. Restlessness at night

due to irritation may be indicative of northern fowl mite infestation

(Petrak, 1969), but again, positive determination of infestation can

best be made by examination of birds.

Latent effects of northern fowl mite infestation

Death of the bird host is often associated with severe northern fowl

mite infestations and could be termed the utmost patent effect. Death,

however, is due to the results of certain latent effects. Cameron

(1938) blamed loss of vitality and death on loss of blood. Although it

is not known whether blood loss produced an anemia, death in severe

cases has been attributed to anemia which resulted from exsanguination

(Metcalf et al., 1962; Petrak, 1969; Koehler, 1977; Matthysse et al.,

1974). Recent studies have shown that this is not necessarily the case.

Loomis et al. (1970) worked with hens having mite populations from light

to severe and anemia was not shown to be a symptom of heavy mite infes-

tations. DeVaney et al. (1977) found no anemia in roosters due to mite

populations.

Weight loss has also been attributed to severe northern fowl mite

infestation (Anonymous, 1967; Koehler, 1977). DeVaney et al. (1977)

found no significant differences in the weights of roosters due to mite

populations. In another study weights of two groups of hens did not

change significantly due to mite infestations (DeVaney, 1979).








One of the longstanding economic reasons for keeping flocks free

of northern fowl mites has been that mites cause a drop in egg produc-

tion (Cameron, 1938; Metcalf et al., 1962; Anonymous, 1967; Koehler,

1977; Rock, 1978; Smith, 1978). Combs et al. (1976)-demonstrated

that chemical removal of mites improved egg production. Other work

done in the last 10 years also conflicts with studies attributing

decreased egg production to northern fowl mites. Loomis et al. (1970)

could find no significant difference in egg production due to mite

populations. Bramhall (1972) discounts northern fowl mites as a

reason for reduced egg production and suggests that poultrymen control

mites only to prevent discomfort to workers. Eleazer (1978) found that

uncontrolled northern fowl mite infestations did not cause reduced egg

production and DeVaney (1979) reported that during two separate 1-year

trials a significant reduction in egg production was produced by mites

for only 1 month in one trial, and 2 months in the other.

Medical importance of Northern Fowl Mites

Allergic reactions

Gamasoidosis, a poultry handlers' dermatitis caused by fowl mites,

was reported in 1824 (Toomey, 1921). It has since been well established

that northern fowl mites will attack man and produce transitory rashes

on the skin (Riley and Johannsen, 1915; Van Der Hoeden, 1964; Frazier,

1969; James and Harwood, 1969; Georgi, 1974; Ebeling, 1975).

Riley and Johannsen (1915) called the mite-produced rash a pruritis

and not a dermatitis since man does not present favorable conditions for

mite viability. Both terms, pruritis and dermatitis, have been used by

recent authors to describe the condition (Cahn and Shechter, 1958;








McGinnis, 1959; Genest, 1960). Papular, vesiculo-papular, urticarial,

or a combination of these primary lesions will develop at the bite site,

the extent and severity of which is thought to be due to an allergic

mechanism (Frazier, 1969). Contact with living miteS may not be

necessary to produce symptoms as both body parts and excretory products

of the mites have inherent toxic properties (Chandler, 1949).

Several non-poultry-related cases of northern fowl mite dermatitis

have resulted from mites entering buildings via window air conditioners

(Cahn and Shechter, 1958; McGinnis, 1959; Genest, 1960). In all of the

cases, abandoned birds' nests were found in or near air conditioner

air intakes. Affected persons were advised to remove mites by bathing

after which all symptomology disappeared in 24 hours. Fumigation of the

buildings and air conditioners, and removal of birds' nests from the

air conditioners eliminated the mite populations.

Disease transmission

When it was found that the chicken mite, Dermanyssus gallinae,

could transmit the virus of St. Louis encephalitis directly and trans-

ovarily (Smith et al., 1945, 1946, 1947), the question arose as to

whether or not the northern fowl mite possessed the same capability.

Collections of northern fowl mites from wild birds yielded mixtures of

viruses containing not only St. Louis encephalitis virus, but also the

virus of western equine encephalitis (Reeves et al., 1947; Hammon et al.,

1948; Bisseru, 1967). The importance of the mite as a vector or reser-

voir for either virus later proved questionable (Reeves et al., 1955).

Subsequent studies have shown the northern fowl mite to be a very poor

transmitter of western equine encephalitis (Chamberlain and Sikes, 1955)








and eliminated it as a possible transmitter of St. Louis encephalitis

(Chamberlain et al., 1957; Chamberlain, 1968).

The northern fowl mite has also been accused of transmitting fowl

pox (Brody, 1936), Newcastle virus (Hofstad, 1949), Lankesterella corvi,

a blood parasite of rooks (Baker et al., 1959), a Bedsonia species of

Ornithosis virus (Meyer and Eddie, 1960), and a microtatobiote, order of

Rickettsiales, of the family Bartonellaceae (Mettler, 1969). Proof of

transmission could not be demonstrated for any of the organisms listed

above.

Control of Northern Fowl Mites

Chemical control has been the method of choice for controlling

northern fowl mites primarily because it is the only method available.

No parasites or predators of the northern fowl mite are known at this

time. Since the mites complete their entire life cycle on the host,

biocontrol agents may not exist.

Many books are available that list various northern fowl mite con-

trols (Hartman, 1953; Benbrook, 1965; Anonymous, 1967; Loomis, 1978).

Benbrook (1965) gives the most comprehensive list of controls prior to

1940, some of which include dust baths containing road dust and wood

ashes, ointments and powders containing mercury compounds, caraway oil

and derris (rotenone), and fumigants such as SO2 and HCN.

Some classes of compounds cannot be used around poultry due to

their toxicity or their tendency to form residues in meat and eggs.

Chlorinated hydrocarbons have been removed from use on or around poultry

due to their formation of residues. Nicotine SOL should be used with

caution since it can be toxic to birds and man. Many organophosphorus









compounds, such as parathion, diazinon, and fenthion (Baytex), have

extremely high avian toxicities and are also excluded from use on or

around poultry (Loomis, 1978).

Little or no research has been done on field application of miti-

cides on poultry. Poultrymen report widespread mite resistance to

labeled miticides, but many of the resistance problems are due to poor

application methods (Eleazer, 1978).

Application methods have changed drastically with the advent of

caged birds and increased flock size. Before 1940, treatment of each

bird in a poultry flock with a dust, ointment, or dip was quite common.

By 1950, the average size of a caged flock was 1500 to 2000 birds

(Hartman, 1953), and the use of treatments that involved the handling

of individual birds rapidly ceased.

A laboratory method was devised for in vitro evaluation of miticides

(Foulk and Matthyssee, 1964). Disposable pipettes are dipped into

miticides and northern fowl mites then drawn inside by use of a vacuum.

The large end of the pipette is covered with fine mesh cloth and after

mites are inside, the small end is plugged with clay. Next, the pipettes

are placed in chambers with controlled temperature and humidity, and

mortality is recorded in 24 hours. This method was also used by Hall

et al. (1978) after slight modification.

Sulfur and nicotine sulfate

These two compounds have been recommended for treatment of northern

fowl mites perhaps longer than others and were initially used because

they had been used successfully for poultry louse control.








The use of sulfur in a dip was recommended by Payne (1929). The

dip consisted of 57 g of sulfur and 28 g of soap for each liter of water.

The dip was only for warm weather use. Emmel (1937) intermittently

fed chickens a diet that was 5% sulfur by weight and.controlled not

only mites, but also fleas and lice. Povar (1946) found that sulfur

actually repelled mites in vitro and the mites continued living for 14

days.

Sulfur has been shown to be effective for northern fowl mite control

when added to poultry litter at the rate of 0.5 kg per 4.7 m2 of litter

(Foulk and Matthysse, 1963). Sulfur is added to poultry litter on the

University of Florida Department of Poultry Science Research Farm and is

routinely used to control northern fowl mites on floor birds (R. H.

Harms, personal communication). A 1% sulfur spray proved ineffective

for northern fowl mite control on caged birds (Furman, 1953).

Nicotine sulfate, or Black Leaf 40TM, has been used as a roost

paint (Payne, 1929; Hansens, 1951), a dust, a dip (Bishopp and Wagner,

1931), and a spray (Povar, 1946; Hartman, 1953). Dips consisted of 1

part 40% nicotine sulfate in 9 parts water with or without the addition

of 28 g of soap per gal of solution (Bishopp and Wagner, 1931). Sprays

contained 1 part nicotine sulfate and 13 parts water. Hartman (1953)

recommended spraying at night and using three treatments at 3-day

intervals.

Nicotine sulfate gave good northern fowl mite control for up to

1 month (Cutright, 1929) and was considered by Povar (1946) to be the

best method of mite control as late as 1946. Furman et al. (1953) re-

ported good, but temporary control with nicotine sulfate. Nicotine








sulfate kills by contact and fumigation. It may cause a 24-hour reduc-

tion in egg production and may kill birds if ventilation is inadequate

(Bishopp and Wagner, 1931).

DDT and lindane

Before their ban due to residue formation, some chlorinated hydro-

carbons were tested on poultry for northern fowl mite control. DDT was

considered an ineffective control when a 10% dust would not control

mites in vivo (Povar, 1946). Lindane (2% EC) gave good results when

sprayed on the vent region of chickens (Hansens, 1951), and a 0.2%

lindane powder is still recommended for northern fowl mite control on

caged exotic birds (Dall et al., 1964).

Malathion

Malathion was at one time an effective compound for northern fowl

mite control. Sprays of 0.25 and 0.5% gave good results at an appli-

cation rate of 25 ml per bird (Hoffman, 1956, 1960). Litter treatments

of 4% dust at a rate of 0.3 to 0.5 kg per 1.9 m2 of litter gave good

results on hens, but severe cases on roosters had to be dusted by hand

(Harding, 1955). In a more recent test, 4 and 6% dusts, and 0.5 and

1.0% sprays of malathion were ineffective for mite control; a 25% dust

gave control for only 3 weeks (Rodriguez and Riehl, 1963). Foulk and

Matthysse (1963) found malathion to be ineffective and suggested that

mites may be showing some resistance to the compound. Perhaps the first

northern fowl mite resistance to malathion in the East was found in

laboratory and field studies by Hall et al. (1978). Nelson and Bertun

(1965) synergized malathion with triethyl trithiophosphate (ethyl DEF)

and increased its toxicity 12.9 times.








In an effort to determine malathion toxicity to fowl, various fowl

were dipped into solutions containing high concentrations of malathion.

A 4% solution killed all birds dipped including one mature goose (Furman

and Weinmann, 1956).

Carbaryl

In the laboratory, 25.0 and 12.5 ppm solutions of carbaryl killed

100% of the northern fowl mites tested (Harrison, 1961). In the field,

a 0.1% solution provided control for only 1 week (Hoffman, 1960). Others

got better results with sprays of 0.25 and 0.5% (Kraemer, 1959; Furman

and Lee, 1969), and 2 to 4% (Foulk and Matthyssee, 1963). Loomis et al.

(1970) got poor control on heavily infested hens with a 0.5% spray, but

Hall et al. (1978) found carbaryl to be the most toxic of the compounds

used in their study.

In tests involving carbaryl dust, Foulk and Matthysse (1963)

achieved good results with a 3% dust but Rodriguez and Riehl (1963)

got control for 22 weeks with a 1% dust.

In studies of the systemic effects of carbaryl on laying hens,

single doses of carbaryl administered orally at 800 and 150 mg per kg of

hen could be detected in the blood for 48 to 72, and 24 hours respec-

tively. Five days after cessation of the smaller dosage, no residues

were found in muscle, liver, fat, skin, or gizzard samples (Furman and

Pieper, 1962). In another study, hens were fed 200 ppm of carbaryl for

7 days. At 3 to 7 days post-treatment, no residues could be found in

muscle, liver, gizzard, skin, or eggs (McCay and Arthur, 1962).

Ronnel

Laboratory and field tests have shown that ronnel is more toxic to

northern fowl mites than either barthrin (a botanical) or malathion








(Bigley et al., 1960). Sprays of 0.25 and 0.5% ronnel effectively con-

trolled field populations of northern fowl mites (Kraemer, 1959; Khan,

1969). Good control was also provided with dusts of 1 and 5% ronnel

(Knapp and Krause, 1960; Foulk and Matthysse, 1963).-

Miscellaneous compounds

Other compounds, mostly organophosphorous compounds, giving good

northern fowl mite control are coumaphos (Bay 21/199) sprays and dusts

(Kraemer, 1959; Hoffman, 1960; Knapp and Krause, 1960; Foulk and

Matthysse, 1963; Khan, 1969), stirofos (SD 8447) sprays and dusts

(Furman and Lee, 1969; Nelson et al., 1969; Combs et al., 1976;

Christensen et al., 1977), dichlorvos sprays and impregnated resin
TM
strands (Khan, 1969; Nelson et al., 1969), crotoxyphos (Ciodrin or

SD 4294) mist sprays (Foulk and Matthysse, 1963), trichlorfon (DyloxTM

sprays (Khan, 1969), naled sprays (Kraemer, 1959), chlordimeform sprays

(Hall et al., 1975; Combs et al., 1976; Christensen et al., 1977), and

neotran and sulphenone sprays and dusts (Furman, 1953; Furman et al.,

1953).

Pyrethroids

Pyrethrum dust has been used with good results on poultry for

northern fowl mite control (Cameron, 1938). Two synthetic pyrethrcids,

EctibanTM and SD 43775, gave good results in laboratory studies. In

the field, effective control was achieved for 57 days with concentra-

tions of SD 43775 ranging from 0.0125 to 0.05% (Hall et al., 1978).

Mechanical controls

The two compounds briefly mentioned here are included only because

they present alternate methods for mite control although the efficacy








or practical value of either one is questionable. Volck, which is

commonly used on plant pests and is nontoxic to birds and mammals, was

tested as an ectoparasite control agent for farm animals (Bruce, 1928;

Caler, 1931). As a 5 to 10% dip, it gave 100% control of northern fowl

mites in 24 hours. Silica aerogel was used to eliminate a northern fowl

mite population that had infested a home via a bird's nest (Tarshis,

1964).

Systemics

Sulfaquinoxaline alone or with other sulfonamides acts as a systemic

acaricide in birds infested with northern fowl mites (Beesley, 1973).

When feed containing 0.033% sulfaquinoxaline was fed to layers, mite

populations were reduced to near zero in approximately 4 weeks (Furman

and Stratton, 1963). Feed with 0.05% sulfaquinoxaline was fed to layers

for 24 hours once a week and an economic control level for northern fowl

mites was reached in 4 weeks (Furman and Stratton, 1964). Out of 15

poultry flocks fed diets containing combinations of sulfaquinoxaline,

sulfadimedine, sulfamerezine, and sulfathiazole for 1.5 to 6 weeks at

concentrations of 0.0125 to 0.02% total sulfonimides, 14 flocks were

free of northern fowl mites at the end of the test (Goldhaft, 1970).












METHODS AND MATERIALS

Laboratory Trials

Environmentally Controlled Rearing Conditions

Rearing of immature diptera was accomplished in a PercivalTM

forced-air, upright growth chamber. The temperature was maintained

at 29.4 C, and continuous lighting was provided by two 40-watt

fluorescent bulbs. The growth chamber was modified to include an

external exhaust system with air supplied to the unit from within the

laboratory. Cages of adult flies of various species were also kept in

this growth chamber unless otherwise specified. Whenever a growth

chamber is mentioned without further clarification in this paper,

reference is being made to the Percival at 29.4 C, continuous lighting,

and ambient humidity lowered by chamber temperature.

Some adult flies were kept in a walk-in growth chamber which had

a relative humidity of 85% and a temperature of 26.7 C. Lighting,

fluorescent and incandescent, was continuous. Since this chamber was

used so infrequently, it will be referred to specifically throughout

the text if applicable.

Colonization and Rearing of Flies

Afsca domestic (L.) -- the house fly. The laboratory house fly

colony was started with adults obtained from a poultry farm in Starke,

Florida, in October of 1975. Wild flies were placed into 3.8-1 plastic

jars half-filled with moistened CSMATM (Consumer Specialties Manufac-

turing Association) and allowed to oviposit. Jars were screened and







placed in the growth chamber. Pupae were removed from the jars,

separated from the CSMA by flotation, and dried on paper towelling.

Care was taken to be sure pupae were free of any mites that may have

been attached to the field-collected adult flies.

Pupae were placed in a standard colony cage (51 X 25 X 25 cm) with

four sides and one end covered with window screen. The remaining end,

fitted with surgical stockinet, was used as an entryway. Water was

provided in a pan ca. 5 cm deep. The water surface was covered with

polyfoam chips to provide a resting area for the flies and reduce

mortality from drowning. The adult diet, a commercially prepared naso-

gastric mixture (Table 5), was thinly spread over a small (ca. 5 X 10 cm)

piece of aluminum foil with the edges raised to resemble a shallow pan.

Additional diet was added daily in thin layers. This method allowed for

a larger feeding surface and reduced waste. Cages with adults were kept

in the walk-in growth chamber.

Eggs were collected over 4-hr time periods in moist CSMA from cages

where females were an average of 7 days old. CSMA was mixed with water

at a ratio of 5:3 and placed loosely in plastic pans 36 cm in diameter

and 14 cm deep. Eggs, about 1000 per pan, were placed 1 to 2 cm below

the CSMA surface to simulate oviposition. Pans were screened and placed

in the growth chamber.

When the colony was well established, a rearing schedule was set up

to provide two cages of flies per week for testing purposes. The

schedule was based on the average time from egg to pupae at 29.4 C

being 10 days. Adults were discarded after 3 weeks.

Interesting contrasts to the above method of rearing house flies

are presented by Grady (1928) and Monroe (1962).







Ophyra aenescens(Wied.) -- the black dump fly. The Ophyra

aenescens laboratory colony originated from adults collected on a west

Florida poultry farm in December of 1976. Eggs were set in pans 25 cm

in diameter and 8 cm deep containing the fortified diet of CSMA, horn

fly dry mix, and water as shown in Table 4 of the results section.

Pupae were separated from the medium by flotation 7 days later, dried,

and placed in a colony cage as for house flies. Besides the water and

nasogastric mix put in the house fly cages (Table 5 of the results

section), adult dump flies were supplied with cane sugar and dry fish

meal.

Eggs were collected in moistened fish meal from 5- to 7-day-old

females. Approximately 500 to 1000 eggs were set twice a week to

maintain the colony.

Hermetia ilZucens (L.) -- the black soldier fly. This fly was

reared in the laboratory on many occasions, but attempts to colonize it

did not succeed. Females placed in jars readily laid eggs on moistened

CSMA or the screened jar lids. Eggs were then set in moistened CSMA as

for house flies. In the growth chamber, larval development required

25 days and pupation another 10 days. Eggs were set primarily to provide

a source of early instar larvae for testing purposes. Besides CSMA,

H. illucens was reared in chicken feed, fish meal, and mixtures of fish

meal and CSMA, all moistened with water.

P::'-O" regina (Meigen) -- the black blow fly. This fly was

attracted to the laboratory during the cooler months of the year and

it was colonized for testing purposes. Females oviposited in moistened

fish meal. Eggs were set in a mixture of 1 part fish meal, 1.5 parts

CSMA, and 1.8 parts water. A medium of fish meal and water was







sufficient for larval development, but the addition of CSMA produced a

lighter textured medium with increased moisture-holding capacity. In

the growth chamber, larval development required 6 to 7 days and pupation

another 5 to 6 days. Adults were maintained on cane sugar, fish meal,

and nasogastric mix as for Ophyra aenescens.

Fannia canicuZaris (L.) -- the little house fly. Fannia was briefly

colonized for a series of experiments. Females would readily oviposit

on the surface of fish meal that was mixed with enough water to make a

semiliquid paste. This mixture was preferred after it aged in the growth

chamber for 24 hr. The surface of the fish meal, which becomes crusty,

could be left in place as the larvae developed, or inverted with the

adhering eggs onto a fresh cup of the fish meal paste. Eggs set weekly

in 120-ml cups of medium provided an ample number of flies. At 29.4 C,

the larval and pupal stages both required ca. 7 days. Adults were

maintained on dry fish meal and cane sugar cubes.

Sarcophaga robusta (Aldrich) -- the flesh fly. While flies were

being reared on fish meal in the growth chamber, sarcophaged flies,

along with other dipteran and coleopteran species, began appearing

inside the growth chamber. This activity ceased when the chamber's

exhaust pipe was covered with a screen. These sarcophagid flies are

also found in poultry manure, so attempts were made to colonize them.

Six females of different sizes were captured and screened inside

360-ml plastic cups with 180 ml of very moist fish meal and placed in

the growth chamber. After females died, they were pinned and labeled

for later comparison with their progeny. Third-instar larvae began

migrating inside the upper halves of the cups 3 days after the females

were screened in. After 2 days of migrating, pupation occurred, and







9 days later, adults began emerging. The size variance in the six

groups of FI adults was greater than the size variance among the six

original females. Microscopic comparisons of the flies, made with

reference to Aldrich (1916) and James (1347), revealed that all

specimens belonged to the same genus and species, Sarcophaga robusta

(Aldrich), syn. S. pZinthopyga (Wied.).

The colony was easily established. Females began mating and larvi-

positing when 5 and 11 days old respectively. Immatures were maintained

as described above and adults were maintained on fish meal and cane

sugar cubes.

Dissection and Mounting of Cephaloskeletons of Third-instar Fly Larvae

Cephaloskeletons of two species of fly larvae were examined for

morphological clues indicative of the modes of life of the larvae.

Third-instar larvae were killed in boiling water and dried on paper

towelling. Each was cut behind the cephaloskeleton so that only a

narrow band of integument still joined the two parts. Next, larvae

were placed in 10% KOH and boiled gently until the unsclerotized

tissues surrounding the cephaloskeletons had dissolved. At the com-

pletion of the KOH treatment, larvae were dried for I-hr periods,

first in 70% and then in 90% ethanol. Larvae were stored in 100%

ethanol. While submerged in 100% ethanol, as much larval integument

and remaining soft tissues as possible were teased from the cephalo-

skeletons. The cephaloskeletons were stored overnight in phenol and

the remaining portions of the larvae were discarded.

Cephaloskeletons were worked into a mixture of phenol-balsam and

placed in desired positions on mounting slides. Care was taken that

the specimens were completely covered with the phenol-balsam mixture.







The mixture was also used to position the glass chips necessary for

coverslip support. At this point, slides were racked and racks placed

in a dust-free enclosure to allow the phenol to evaporate. Three or

four days were necessary for the evaporation step to be completed. This

step can be hastened by placing slides in an oven at 50 C for 48 hr. If

coverslips are added before phenol has completely evaporated, specimens

may be damaged.

After the phenol had evaporated, coverslips were placed over the

specimens using pure balsam. Slides were set aside until the balsam

had dried.

Bioassay of Poultry Manure

Manure for laboratory bioassay was collected in 360-mi plastic

cups. Samples were capped and then frozen for a minimum of 24 hr to

kill fauna present in the manure. Prior to testing, samples were

removed from the freezer and allowed to thaw completely. Twenty-four

hours were usually required for samples to reach ambient temperature.

Unless otherwise specified, manure samples were seeded with first-

instar larvae of the particular fly species to be tested. Eggs were

used exclusively at first but their use was discontinued when first-

instar larvae produced more precise results. Manure was never recon-

stituted with water.

After larvae were added, samples were covered with screen and

placed in the growth chamber. Adults were allowed to emerge and die

prior to examination of samples. Pupae and adults were separated


from manure by flotation.







Addition of a Liquid Insect Growth Regulator (IGR) to Larval Media of
Flies

In order to simulate conditions in the field, larval diets were

moistened with water containing various levels of a liquid IGR.

Control diets were prepared using plain water.

Diets were placed in 360-mi cups and first-instar house fly larvae

were added instead of eggs. Cups were covered with screen and put in

the growth chamber until pupae closed and adults died.

Laboratory Tests with Granular Baits

Knockdown tests. Test baits were sprinkled in brown paper bags,

21 by 13 by 6 cm, which had been stapled side by side to a piece of

wood ca. 61 cm in length. This arrangement of baits was stored outside,

under the eaves of the laboratory, to simulate actual weathering

conditions.

On the morning that baits were placed in the bags, the knockdown

test was conducted. Three- to five-day-old female colony house flies

were transferred by means of a vacuum system to cylindrical window

screen cages, 12 cm high by 7 cm in diameter. Cages were inverted over

the baits with 10 flies per cage and four cages per bait. The cages

had no bottom surface and allowed flies to come in direct contact with

the baits. Mortality was noted at 10-min intervals throughout the day

until all flies had died. Criterion for death was total lack of move-

ment. After the test, baits were stored as described above.

Residual tests. At some time interval after the knockdown test

and at selected intervals thereafter, the residual activity of the same

bait samples used above was tested until daily fly mortality was less

than 50%. Flies were exposed to the baits as described above and

mortality was recorded after a 6-hr exposure period.








Attractiveness tests. Baits were sprinkled into bait stations

fashioned of 3.8-1 milk jugs (R. C. Axtell, unpublished data). Bait

stations were placed in a 1.8 by 1.8 by 3.7 m screened enclosure into

which 200 five-day-old female house flies had been released. The

enclosure was in full sun but baits were protected from sun and rain

by a small structure inside the enclosure. Mortality was recorded

after 24 hr.

Topical Application of Insecticides to House Fly Adults

Stock solutions were made by placing 1 g of the active ingredients

(Al) of the insecticide in 100-ml volumetric flasks and adding enough

acetone to bring the volumes up to 100 ml. Test concentrations were

made in acetone from serial dilutions of the stock solutions.

Before use, all glassware was washed in a detergent and rinsed

thoroughly with water. When dry, three final rinses of acetone were

applied and glassware was baked in an oven at 176.7 C for 24 hr.

Laboratory colony house flies, 3 to 5 days old, were immobilized

with a vacuum and males discarded. While immobilized, female flies
TM
were dosed with test concentrations using a 10-pl HamiltonTM syringe

equipped with a HamiltonTM repeating dispenser. Flies were released

into cylindrical cages, 12 cm high by 7 cm in diameter. Cotton balls

saturated with a sucrose solution were placed on the tops of cages as

a food source.

Tests were performed at 26.1 C and ambient humidity. Mortality

was recorded after 24 hr. Criterion for death was total cessation of

movement.








Laboratory Bioassay of Acaricides

Northern fowl mites were exposed to various dosage levels of

acaricides to collect data necessary for dosage-mortality curves.

The testing procedures were adapted from those of Hall et al. (1978).

Tests were standardized as suggested by Peet and Grady (1928).

Squares of muslin cloth were secured with neoprene bands over

the wide ends of 23-cm disposable Pasteur pipets. Acaricides were

dissolved in acetone and serially diluted with acetone to the desired

concentrations in final volumes of 100 ml. Pipets, with cloth squares

in place, were immersed in the acaricide solutions for ca. 20 sec, then

removed and rolled on paper towelling to dry the outsides. Control

pipets were treated with 100 ml of acetone. Next, pipets were tapped

on paper towelling for 2 min, tapered ends down, to dry the insides.

More complete drying was achieved by using a GastTM electric pump to

force air through the pipets for 20 min. Pipets were removed from the

pump and used within I hr.

After pipets were ready for use, mites were collected from caged

chickens at the University of Florida Poultry Science Farm and trans-

ported to the laboratory. Mites were emptied into a porcelain emesis

basin which was placed inside a larger pan half-filled with water to

prevent mites from escaping. The vacuum side of the above-mentioned

pump was fitted with a length of neoprene tubing and the pressure set

at 106 g/cm2. The large ends of the pipets, with cloth squares attached,

were slipped into the open end of the neoprene tubing. When the pump

was turned on, mites were pulled into the pipets. After the desired

number of mites were inside, the pipets were removed from the tubing.

The tapered ends were snipped with a hemostat to a length that would







allow the pipets to stand diagonally inside 1000-ml beakers and the

tips were sealed with clay.

Desiccators were fashioned from 1000-ml beakers. Salt solutions

of 4 g of NaCl and 10 ml of water were added to the beakers to maintain

the relative humidity at approximately 80%. Dry 10-ml beakers were

placed inside the 1000-ml beakers to receive the pipet tips and keep

them out of the salt solutions. Once the sealed pipets were inside,

desiccators were covered with two layers of saran secured with a rubber

band. Desiccators were placed in the growth chamber at 29.4 C.

Mortality was recorded 24 hr later with complete cessation of movement

the criterion for death.

One pipet containing 15 gravid female mites served as a replica-

tion. Each treatment was replicated four times. Control mortality

averaged 11.26% and was never higher than 16.39%.

Field Trials

Rotovation

Rotovation is a term coined by poultrymen in the Tampa, Florida,

area to describe a method of mechanically stirring manure in poultry

houses to keep it dry and unattractive to flies. Manure is composted

in place and can be used for fertilizer without further treatment.

The tilling unit, made by Selpats Manufacturing, Inc., P. 0. Box
TM
149, Palatka, Florida, 32077, is officially called the Dryovator .

The tiller is operated by the power take-off of a modified Kuboda L175

diesel tractor. Tractor and tiller are shown in Figure 1.

The act of rotovating is termed rototilling or more frequently,

just tilling. Tilling was accomplished by driving the tractor down

the walkway of a poultry house and pulling the tiller through the












































Figure 1.


View of tractor and tiller.







manure on either side of the walkway. Houses were tilled by pulling

the tiller down one walkway and up the other. The process was reversed

each time a house was tilled in order to more thoroughly stir the

manure. This became the standard procedure in all tilling trials, even

when houses were tilled less frequently than once a day. Figure 2

shows the tiller in operation. Since the dimensions of poultry houses

vary from farm to farm, tillers must be custom-made for each farm.

Description of the Tilling Site

Tilling trials were accomplished on a north Florida poultry farm

near Starke, Bradford County. Prior to construction of the farm,

earth was removed so that the foundations of the poultry houses were

0.9 m lower than the level of the ground immediately surrounding the

farm. This complicated drainage problems, especially during periods

of wet weather. The farm consisted of four California-style flat-deck

houses (Figure 5) 90 m in length containing 5000 chickens each, and

one double-wide stair-step house containing 15000 chickens. All birds

were housed three to a cage. Only the California-style houses were

used in the pest management studies. The layout of the farm and the

designation of the houses are shown in Figure 3. The watering system
TM
consisted of one HartTM cup per every two cages. This system worked

well when properly maintained and cups were routinely cleaned. Water

and feed were free choice.

Monitoring Larval Fly Populations

Techniques developed for field evaluations of larval fly popula-

tions included the use of pupal traps. Cylinders 31 cm tall by 10 cm

in diameter made of 1-cm mesh hardware cloth were filled with moistened

wood chips and inserted into the chicken manure pack in poultry houses.






































a :.f- ,-.~
r~~a .
'~-ri I- ~i
I tr~a


Figure 2. Tiller in operation.











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A golf course turf plugger 10 cm in diameter was used to make the holes

in which the cylinders were inserted. These cylinders, or pupal traps,

were highly attractive to third-instar larvae as pupation sites and

could therefore be used to collect fly pupae of known age.

In placement of pupal traps, areas in the manure pack were selected

that appeared suitable for fly production. Once a site was chosen, the

plugger was used to remove a horizontal plug of manure from the edge of

the manure pack. The pupal trap was placed in the resulting hole and

firmed into place. A tag with identifying data was tied to the bottom

of the chicken cage directly above the trap to aid in locating the trap

at a later date.

Collecting the traps was simple once they were located. Layers of

manure made traps difficult to find at times even with the aid of the

tags. Once found, traps were removed from the manure pack and tagged

(Figure 4). Plastic bags are advisable for transport of traps after use.

Pupae were separated from the wood shavings by flotation. Trap

contents were emptied into suitable containers and water was added.

After ca. 30 min, wood shavings sink leaving only the pupae floating on

the surface. If enough containers were available, all trap contents

were floated simultaneously.

Poultry and Poultry Facilities Used When Evaluating IGR's as Oral
Larvicides

When IGR's were tested as oral larvicides, the amount of manure

needed for sampling and the frequency with which it was collected deter-

mined the number of hens used per treatment group. A hen voids ca. 92 ml

of wet manure per day or 647 ml per week (Hart, 1963). Ten hens will

produce 6468 ml of manure weekly, which is enough to provide a maximum












































A tagged pupal trap after removal from
manure pack.


Figure 4.







of five 360-mi samples three times per week for laboratory bioassay.

Therefore, the minimum number of hens used in a treatment group was 10.

To consolidate manure as much as possible, hens were housed two to

a cage in cages measuring 20 by 45 by 43 cm. To prevent cross-contami-

nation of manure, two cages were left empty between treatment groups

and/or vertical tin dividers ca. 46 cm high were placed in the manure

collection area between treatment groups. Manure collection areas were

cleaned out before experiments began and covered with sheets of poly-

ethylene or tin to catch the treated manure.

When IGR's were mixed with feed, vertical dividers were placed

between feed troughs of treatment groups to prevent hens from sampling

treatments other than their own. The continuous watering troughs used

by all treatment groups were directly below the feed troughs and cross-

contamination was possible via spilled feed. To minimize this problem,

clay dividers were placed in the water troughs and water was indepen-

dently piped into and drained out of the sections of trough that served

each treatment group.

When IGR's were mixed with drinking water, water troughs were again

divided between treatment groups. Water treatments were given to the

chickens at 9:00 a.m., 12 noon, and 4:00 p.m. daily in the amounts of

50 ml per bird per treatment. Care was taken not to spill treated water

into manure collection areas. Water troughs were lined with poly-

ethylene to prevent them from being contaminated by unlabeled compounds.

Feed, treated or untreated, was always offered free choice and

water was provided in either a continuous gravity flow system or on the

schedule described above. Hens were used instead of roosters so that

egg production could be monitored if desired. To maintain a maximum







rate of lay, hens were exposed to 14 hr of light by the use of supple-

mental incandescent lights in the morning and evening. Ail eggs

produced by hens consuming unlabeled IGR's were destroyed and hens were

destroyed after the experiments were terminated.

Calculation of Hen-Day Egg Production and Average Daily Feed Consumption

The term hen-day implies that during the time period over which the

calculations for production or consumption have been made, daily hen

mortality has been taken into consideration. In order to calculate on

a hen-day basis, daily mortality records were kept.

Hen-days are calculated by multiplying the number of hens on hand

by the number of days in the designated time period. This is simple if

no mortality occurs. For example, 10 hens during a 7-day period consti-

tute 70 hen-days. However, if one hen died on day 5, the number of hen-

days becomes nine hens times 7 days plus one hen times 4 days for a total

of 67 hen-days. Whether or not the day on which hen mortality occurs is

to be counted in the calculations for hen-days should be decided in

advance and adhered to.

Hen-day production is therefore the number of eggs laid during a

time period divided by the number of hen-days calculated for the same

time period. The quotient is multiplied by 100 since hen-day production

is expressed as a per cent.

Consumption figures are usually expressed as the amount of feed

consumed per bird per time period, e.g. 114 g per bird per day, and

not as hen-day consumption. However, total hen-days as well as total

feed consumption must be known in order to calculate the average daily

consumption per bird. Total feed consumption is all the feed consumed








over a time period, which is found by weighing feed at the beginning

and end of the designated period and subtracting the two figures.

Average daily consumption is therefore the total feed consumed

during a time period divided by the hen-days for the time period.

Addition of IGR's to Poultry Feed

Insect growth regulators (IGR's) were added to the University of

Florida Basal Layer Diet (Table 1). The basal diet was mixed in
TM
136.4-kg lots in a Kelley-DuplexTM vertical mixer. When the basal diet

was thoroughly mixed, ca. 4.5 kg were removed to a small paddle mixer

where the appropriate amount of IGR was added while the mixer was in

operation. This premixture was mixed for 10 min, returned to the

vertical mixer, and added slowly to the mixing basal diet. After all

of the premixture was added, mixing continued for 10 min. Diets were

removed from the vertical mixer in six 22.7-kg batches and placed in

aluminum cans for ease of handling. Feed cans were labeled and dated

for identification purposes.

The vertical mixer was cleaned by swirling 11 kg of cracked corn

inside the mixer for 10 min. The corn was removed and discarded. Next,

fine feed particles were removed from the mixer with compressed air.

The paddle mixer was cleaned with a whisk broom and compressed air.

Mixers were always cleaned between mixes.

Topical Application of Granular IGR's to Poultry Manure

In the manure collection area of one poultry house at the tilling

site, a granular IGR was applied to manure with a hand-held fertilizer

spreader. Granules were preweighed at the laboratory and the amount

for each treatment was individually placed in the spreader. The IGR

was applied uniformly to manure treatment blocks until the spreader

was empty.








Table 1. Composition of basal diet for poultry feed trials.


Ingredients


Per cent


Yellow corn

Soybean meal (50% protein)

Alfalfa meal (20% protein)

Ground limestone

Dicalcium phosphate

Iodized salt


69.33

19.00

2.50

6.17

2.25


Micro-ingredient mixa .50

Total 100.00

C.M.E./kgb 2890

Per cent protein 16.2

Per cent calcium 2.98

Per cent phosphorus .73


a Supplies per kg of diet; 6600 I.U. vitamin A; 2200 I.C.U. vitamin
DA; 11 I.U. vitamin E; 2.2 mg menadione dimethyipyrimidinal bisulfite
( PB); 4.4 mg riboflavin; 13.2 mg pantothenic acid; 59.6 mg niacin;
998.8 mg choline chloride; 22 mcg vitamin 812; 110 mcg biotin; 125 mg
ethoxyquin; 60 mg manganese; 50 mg iron, 6 mg copper; 0.198 mg cobalt;
1.1 mg iodine; 60 mg zinc.


Calories metabolizable energy per kilogram.
Calculated values.
Calculated values.








On the University of Florida Poultry Science Farm, granules were

uniformly applied to manure collection areas beneath treatment blocks

of cages with a shaker fashioned from a 480-mi glass jar. Amounts were

preweighed and applied to manure treatment blocks until the shaker was

empty.

Mixing and Application of Liquid IGR's and Organophosphorus Larvicides

Liquid IGR's and commercial larvicides were applied to manure with

a 7.7-1 SearsTM pressure sprayer. The nozzle was adjusted so that the

spray was emitted in a broad cone. Each treatment was applied to its

block by computing the volume of larvicide to be applied, mixing the

volume in the sprayer, and applying the volume uniformly to the par-

ticular treatment block until the sprayer was empty. The sprayer was

cleaned thoroughly with water between applications of treatments.

Samples consisted of four 360-mi cups of manure collected from the

center third of each treatment block. After a sample was collected, it

was emptied into an aluminum pie pan. The pan was placed in the sun and

the living larvae of selected fly species present in the sample were

counted. Criterion for death was total cessation of movement.

Addition of a Liquid IGR to the Drinking Water of Hens

The facilities and water treatment application techniques used were

described in the poultry facilities section. Test concentrations were

prepared by serially diluting IGR stock solutions of 0.1 and 1.0%.

Samples were bioassayed in the lab using first-instar house fly larvae.

Placement of Light Traps

Two blacklight electrocutor grid traps were evaluated at the tilling

site. One trap was hung in the aisle between house 3 and the egg pro-

cessing room, and the second was hung in the same aisle, but between








house 4 and the egg cooler (Figure 3). Traps were 1.8 m above the

ground and 6.i m apart. Both traps were similar in design but the one

near the egg processing room, trap A, was yellow and the other one,

trap B, was black (Figures 5 and 6). The manufacturer stated that the

light sources for the two traps were producing light at different wave

lengths, but exact values were not disclosed.

The traps were automatically turned on along with the farm's

supplementary lighting system in the morning. They were in operation

all day and were turned off at night along with the farm's supplemental

lighting system. This reduced the collection of insects other than

those associated with the poultry farm, e.g. nocturnal moths.

Traps were emptied weekly and the contents transported to the

laboratory in plastic bags. Bags were labeled and frozen until contents

could be analyzed. Catches were analyzed by counting selected species

of flies in a representative sample of each catch. Samples consisted

of a volume of each catch that weighed 10% of the total catch weight.

The number of flies in a catch was assumed to be the number of flies

counted in the sample multiplied by 10.

Field Tests with Granular Baits

Bait stations were fashioned from brown paper bags, 21 by 13 by 6 cm.

The 6-cm lip helped keep baits and dead flies from being blown from the

bait station by strong winds.

When testing was done at the tilling site, bait stations were

placed at the sun-shade interface on the south sides of the poultry

houses and secured by punching a 12-penny nail through the bag and into

the ground. Baits were added to the bags after bags were secured.































I .


Ili
* I.I !
* .;*


Light trap opposite egg processing room.
Note the flat-deck cage arrangement.


Figure 5.



















































Figure 6. Light trap between egg cooler and
house 4.








Following a 6-hr exposure period, each bait and station was placed

into a plastic bag, returned to the laboratory, and the catch processed

by sex.

When testing was done at the University of Florida Thoroughbred

Unit at Ocala, Florida, bait stations were spaced along the edge of the

concrete center aisle of a horse barn and each was secured with a rock.

After a 24-hr exposure period, baits and stations were collected

and processed as above.

Application of Contact Residuals to Selected Surfaces

Templates of plywood, cement block, and galvinized tin were selected

for use in residual tests because these are the types of surfaces most

likely to be sprayed with a contact residual in poultry houses.

All templates were cleaned with soap and water and allowed to air

dry prior to treatment. Pesticides were applied to run-off with a hand-

held trigger-action sprayer. The first test began as soon as the

templates had dried.

Another method for testing residuals by use of blotting paper

templates is described by Batth (1974).

Application of Contact Residuals to Plywood Panels

Panels, 61 by 122 by 0.6 cm, were cut from 1.2 by 2.4 m plywood

sheets and designed to hang with the long side in a horizontal position.

Panels were hung by attaching two 46-cm lengths of light-weight chain

to the upper corners. Aluminum rain gutters, for catching insects

killed while on the panels, were placed horizontally along both sides

of each panel so that the bottom of the guttering was even with the

lower edge of the panel (Figure 7).











































Panel with guttering suspended by chains
at the tilling site.


Figure 7.







Compounds were mixed using formulas as described in Neal (1974)

and applied to run-off with hand-held trigger-action sprayers. Nozzles

were adjusted to produce a cone 15 to 20 cm in diameter when the sprayers

were held 31 cm from the panel surface. Sprayers were calibrated with

graduated cylinders.

After insecticides were applied and allowed to dry, panels were

hung in houses 1 through 4 at the tilling site, and the guttering was

attached.

Evaluation of Northern Fowl Mite Populations

Field estimates. Field evaluation of mite populations on individual

birds required two workers. The first worker, the handler, suspended the

birds by their feet with the birds' breasts facing the second worker, the

counter (Figures 8 and 9). The counter examined the birds, starting at

the tip of the keel bone, working caudally to the vent area, and over

the dorsal portion of the tail. In severe cases, mites were found on

both legs down to the shanks, and more anteriorly than the tip of the

keel bone.

Counts were designated as follows:

No mites seen 0
From 1 to 10 mites Counted individually
From 10 to 100 mites Counted by 5's, i.e. 15,20,25,etc.
From 100 to 200 mites Counted by 50's
Over 200 mites Counted by 100's

Counters and handlers never interchanged. Counters identified the

birds and recorded the mite counts after birds were examined. Counters

frequently double-checked each other to be sure that counts were uniform.

Calculation of a conversion factor. An attempt was made to correlate

field-estimated mite populations with mite populations actually present

on hens by extracting field-estimated mite populations from hens with a
































A pair of workers examining a hen for mites.
The counter is on the left. Note the
stair-step cage arrangement.


A close-up view of Figure 8.
area on the chicken is due to
mite debris.


The darkened
mites and


Figure 8.


Figure 9.








soap and water solution. Ten birds with five different levels of field-

estimated mite populations were washed and the mites counted in the

laboratory. The field estimations, the laboratory estimations, the

ratios between the two, and the mean of the ratios are shown below:

Field Laboratory Lab Est.
Estimation Estimation Field Est.

100 2710 27.10
500 2895 5.79
1000 5065 5.07
2500 4735 1.90
5000 4050 0.81
x = 8.13

The mean of the ratios between the laboratory estimate and the

field estimate was used as the correction factor. Field estimates

were multiplied by 8.13 to arrive at a corrected field estimate.

Unless otherwise stated, mite values referred to in the text are

field-estimated values. Converted values are for reference only.

Field Application of Acaricides to Caged Hens
TM
Acaricides were applied to caged hens with B.& G. cans and a

Sears 7.7-I pressure sprayer. Nozzles on both types of sprayers were

adjusted to emit cones ca. 15 cm wide when nozzles were held 31 to

46 cm below the cages. Acaricides were applied from beneath the cages

in an effort to thoroughly soak the vent areas of the chickens. When-

ever possible, applicators stood on the side of the cages opposite the

feed trough to give then an unobstructed view of the hens while applying

the acaricides.

Acaricides were mixed and applied according to directions found

in the Insect Control Guide (FAES). Sprayers were cleaned thoroughly


between treatments.








Field Application of Acaricides to Floor Birds

Birds were suspended by their feet and sprayed individually. The

area between the keel bone and the tail was thoroughly saturated with

the acaricide solution. Application was made with a B.& G. can. Litter

was not treated.

Compounds Utilized for Fly or Mite Control

Common names, code letters and numbers and/or trademarks of the

compounds utilized in this study are shown in Table 2. Names which are

in accordance with the principles of Chemical Abstracts nomenclature

are given if available (Kenaga and End, 1974). If the compound was

supplied by a cooperator, the manufacturer's name is included. Compounds

without a manufacturer's name were purchased locally.

Treatment of Data

Statistical Analysis System (SAS). Data were analyzed at the

Northeast Regional Data Center (NERDL), University of Florida, Gaines-

ville, Florida, using the Statistical Analysis System (SAS) of Barr

et al. (1976, 1979).

Comparison of means. Methods for comparison of means, such as chi

square and Tukey, were found in Snedecor (1961) and Freese (1963).

Duncan's multiple range tests were performed by SAS.

Probit analysis. Probit analysis and the plotting of dosage-

mortality curves were performed by SAS. An in-depth explanation of

probit analysis and the calculation of a probit line was found in

Finney (1964). Aid in interpretation of probit lines was given by

Hoskins and Gordon (1956) and Tsukamoto (1963).

Correction of mortality. Where applicable, the results of pesti-

cide trials were corrected by the methods of Abbott (1925).
















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RESULTS

House Flies

Manure Management

Tilling wet manure. At the tilling site, fairly dry manure, 10 to

15 cm deep, had become wet from seasonal blowing rains and threatened

to overflow onto the walkways. An attempt was made to dry the manure

by tilling every house twice a day, 7 days a week. Houses 1 through 4

were tilled and then the process was repeated after a 30-min interval.

Tilling was done during the noon hour when the temperature was high and

workers were not in the houses.

Results. When tilling began, the manure pack was not uniformly

wet, but too wet in most places for house flies to breed. The presence

of house fly adults was hardly noticed and soldier flies, if present,

were not evident. Manure was a shapeless mass with the consistency of

a thick paste. Problems were compounded in some areas by leaking

Hart cups.

Tilling tended to dry and texturize as well as push manure away

from the walks and leave it in mounds towards the center of the manure

collection areas (Figure 10). After 1 week of tilling, the manure pack

began to hold its shape, but flowed back to the edges of the walkways

after 24 hr. Although the moisture level had dropped by a noticeable

amount, the manure had the consistency of mashed potatoes and was not

yet breaking into individual pieces when tilled.

















































Figure 10.


The appearance of fairly dry manure after
tilling. Note how manure is pushed away
from the walk and mounded in the center
of the manure collection area.







As drying increased, pockets of house fly larvae began to show up

in areas now suitable for their development. Tilling stirred the flies

and caused them to reorient at the manure surface, but it is doubtful

that tilling at this rate prevented them from completing their cycles.

Pockets of maggots tilled one day had reformed by the next. Soldier

flies were still not present in large numbers and the manure was now

becoming drier than they preferred.

By the end of the second week, manure began to break up into chunks

ranging from 3 to 10 cm in size (Figure 11). Although this was a sign

that the moisture level of the manure was decreasing, numerous pockets

of house fly larvae were proof that the manure was still not dry enough

to retard their development. The manure pack now held its shape over-

night and no longer threatened to overflow onto the walkways.

By the end of the third week, manure was becoming more friable.

In most areas, manure had broken into 3-to 5-cm chunks which were

crusty on the outside and wet on the inside. Drying continued and

pockets of house fly larvae became fewer in number. The manure pack

was gradually losing volume due to the drying process. This was evident

from the increased space in the manure collection area, i.e. the space,

after tilling, between the walk and the manure pack, and by the decrease

in the amount of manure thrown onto the walks while the tiller was in

operation.

At this time, the farm owner decided that the manure was dry enough

to be removed from the houses. Despite my suggestions that he wait

until a later date, the manure was removed and the tilling program

terminated.































Figure 11.


Manure which has dried enough to form
particles of various sizes when tilled.


^1"0







No rain fell during this tilling experiment. Temperatures were

between 29.4 and 32.2 C during the day and a stiff breeze was blowing

at ground level.

Results recorded during this and other tilling experiments were

mostly subjective due to the difficulty in utilizing objective sampling

methods. Pupal traps could not be employed because of the tilling

schedules and facilities for drying manure were not available when all

tilling experiments were performed.

Tilling after the addition of wood chips and sand to manure. When

manure had completely liquified due to blowing rains, and tilling was

ineffective, builder's sand and wood chips were added to manure in

houses 1, 2, and 4 to improve the consistency. The experimental design

is shown in Figure 12. The manure collection areas between the walks

were treated and evaluated. The treatment blocks in houses 2 and 4 were

7.44 m2. House 1 was divided in half, and chips and sand were put in

the back(A) and front(B) halves respectively (see Figure 12). Chips

were added until they were an average of 5 cm higher than the walk

after spreading. Equivalent amounts of sand were added to the assigned

blocks (Figure 12). House 3 was tilled, but no chips or sand was

added. After spreading chips and sand with rakes, all four houses were

tilled. Figures 13 and 14 show the appearance of the chips before

spreading and after the initial tilling. Tilling continued on a daily

basis for only 11 days, at which time the poultryman decided to clean

out the houses.

Results. Sand was found to be ineffective for improving the con-

sistency of liquified manure. It was heavy and difficult to distribute

in the houses. When moistened by the manure, the sand became even






























Figure 12.


1
4-N

C





S


2



S
B
C
B
S
B
Cr
B
iS


3




C
0
N
T
R
0
L


4

-1
B
C
B
S
B
C
B
S
B
C


C=CH!PS
S=SAND
B=B LANK


Experimental design for adding builder's sand and wood
chips to houses 1 through 4 at the tilling site.
Only the front two-thirds of the houses are shown.


































The appearance of chips before spreading.


S_. ?


r.- ;. .- r." ...
.-, ;. ;-< -* *



i--~r ~C.13' -^*:)' r
i.- :..: ..,.-:.. :
.; ,: : .--
..A
k .. r ".Z .., ',..


Figure 14.


The appearance of chips after the initial
tilling.


Ar,



-r





ye*
. .~



O" 9P
,,% .A


Figure 13.







heavier and made the manure difficult to remove when poultry houses were

cleaned out. The extra weight of the sand could not be tolerated by the

manure spreader owned by the poultryman, and he voiced his dissatisfaction

after having to make several minor repairs as a result.

Instead of aerating manure, sand packed it down tight. Sufficient

amounts could not be added to wet manure to provide the consistency

needed for tilling without causing the manure to be too heavy.

Wood chips proved to be an excellent additive to liquified poultry

manure. Chips were light and easy to handle. They aerated and aided in

drying manure, and facilitated manure removal. Wood chips were also 50%

cheaper in price than sand and more readily available.

After chips had been added and manure was tilled once a day for 2

days, the manure had a consistency that was still wet, but friable.

Fresh manure had a relatively dry bed to fall upon before being tilled.

Chips did not pack like sand, but remained light and enhanced drying by

providing increased surface area.

By the 11th day, the areas where chips had been added were still

wet but in much better condition for removal from the poultry houses

than was the manure in other treatment groups. The control house was

unchanged and the manure had the consistency of thick soup. The houses

where sand had been added were essentially the same as the control, but

some areas now had a thicker, heavier consistency.

No rain fell while the experiment was performed, but skies remained

overcast. Temperatures averaged 27 C and the air was calm.

Tilling with and without the addition of wood chips to manure. On

one occasion when the poultryman had his houses cleaned out, the manure

and the sand beneath it, both of which were dry, were removed to a level