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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
Added title page title:
Poultry pest management techniques in Florida poultry houses
Creator:
Hogsette, J. A ( Jerome Adkins ), 1945-
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Language:
English
Physical Description:
xvii, 308 leaves : ill. ; 28 cm.

Subjects

Subjects / Keywords:
Birds ( jstor )
Chickens ( jstor )
Hens ( jstor )
Larvae ( jstor )
Manure ( jstor )
Mites ( jstor )
Mortality ( jstor )
Poultry ( jstor )
Pupae ( jstor )
Tillage ( jstor )
Housefly -- Control ( lcsh )
Mites -- Control ( lcsh )
Poultry -- Diseases -- Control ( lcsh )
Poultry industry -- Health aspects -- Florida ( lcsh )
City of Chipley ( local )
Genre:
bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )

Notes

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

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University of Florida
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University of Florida
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Copyright [name of dissertation author]. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
Resource Identifier:
020715705 ( ALEPH )
AAB7411 ( NOTIS )
06300526 ( OCLC )

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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
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L


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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.- :..: ..,.-:.. :
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Figure 14.


The appearance of chips after the initial
tilling.


Ar,



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ye*
. .~



O" 9P
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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




Full Text

THE EVALUATION OF POULTRY PEST MANAGEMENT
TECHNIQUES IN FLORIDA POULTRY HOUSES
BY
JEROME ADKINS HÃœGSETTE, JR.
A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL
OF THE UNIVERSITY OF FLORIDA IN
PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR
THE DEGREE OF DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
137?

ACKNOWLEDGEMENTS
I would like to express my gratitude to Dr. Jerry F. Butier 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.
! 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 Chip ley, 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 il
LIST OF TABLES vi
LIST OF FIGURES xi i
ABSTRACT xv
INTRODUCTION 1
LITERATURE REVIEW
The House Fly
History and Economic Importance ,
B ionom i cs
Methods for Larval Control
Methods for Adult Control in Poultry Houses
Northern Fowl Mites
Description, Biology, and Control
Effects on the Host
Medical Importance of Northern Fowl Mites..
Control of Northern Fowl Mites
3
3
4
7
21
27
27
32
34
36
METHODS AND MATERIALS
Laboratory Trials
Environmentally Controlled Rearing Conditions
Colonization and Rearing of Flies.....
Dissection and Mounting of Cephaioske1etons of
Third-Instar Fly Larvae
Sioassay of Poultry Manure
Addition of a Liquid Insect Growth Regulator (IGR)
to Larval Media of Flies
Laboratory Tests with Granular Baits
Topical Application of Insecticides to House Fly Adults
Laboratory Bioassay of Acari cides
Field Trials
Rotovat icn
Description of the Tilling Site
Monitoring Larval Fly Populations
Poultry and Poultry Facilities used when Evaluating
IGR's as Oral Larvicides
Calculation of Hen-Day Production and Average Daily
Feed Consumption
Addition of IGR's to Poultry Feed
43
43
43
47
48
ho
4q
50
51
52
52
54
54
57
60
61
i 11

PAGE
Topical Application of Granular IGR's to Poultry Manure 61
Mixing and Application of Liquid IGR's and
Organophospnorus Larvicides 63
Addition of a Liquid 1GR 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 7b
House Flies 76
Manure Management 76
Ophyra aenescens Basic Biology Studies 91
Competition Studies with Eermetia ittueens 1|4
insect Growth Regulators and Grganophcsphorus Larvicides 123
BlacKlight Electrocutor Grid Traps for Adult Fly Surveys 161
Efficacies of Granular Fly Baits 167
Contact Residuals 183
Northern Fowl Mites 1 94
Dosage-Mortality Curves for Selected Acaricides 194
Control of Endemic Florida Strains of Northern Fowl
Mites with Carbaryl , Malathion, and Ravap 193
Efficacy of Two Synthetic Pyrethroid Compounds Against
Northern Fowl Mites on Laying Hens in Floor Pens 2C6
The Effects of Northern Fowl Mites on Egg Production 212
DISCUSSION 239
The Value of Rotovation as a Method of Manure Management 239
Ophyra aenescens Larvae as Predators of ?íusca domestica
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 24°
The Influence of Larvae of Eermetia vVlucens on Other
Species of Fly Larvae 250
The Efficacy of Dimil in as a Feed Additive 251
The Efficacy of Methoprene as a Feed Additive 252
Methoprene as a Topical Larvicide 252
Laboratory Studies with CGA 72662 253
CGA 72662, Dimethoate, Dichlorvos, and Ravap as
Topically Applied Larvicides 253
The Efficacy of CGA 72662 in Water 25,;+
1 v

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 253
The Effects of Northern Fowi Mites on Egg Production 260
Evaluation of the Mite Rating System 262
CONCLUSIONS 264
LITERATURE CITED 267
APPENDICES
1A RAW DATA FROM FIRST OPHYRA AENESCENS ADULT LONGEVITY
STUDY 292
13 RAW DATA FROM SECOND OPHYRA AENESCENS ADULT LONGEVITY
STUDY 293
1C RAW DATA FROM THIRD OPHYRA AENESCENS ADULT LONGEVITY
STUDY 294
ID RAW DATA FROM FOURTH OPHYRA AENESCENS ADULT LONGEVITY
STUDY 296
2 RAW DATA FROM CGA 72662 LABORATORY STUDIES 293
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 212 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 1C I ECTI3ANâ„¢
DOSAGE-MORTALITY DATA 305
8 THE PROBABILITIES, PROBITS, LOG DOSES, AMD 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
308
v
BIOGRAPHICAL SKETCH

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 S3
6. Summary of the four Ophyra aenescens adult longevity studies
including average adult life span and the length of the life
cycie 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 Ophyra 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 Ophyra
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 Musca domestica 103
12. Results of the competition study between Ophyra aenescens
and Musca domestica. 104
13. Experimental design of Ophyra aenescens predation study 106
14. Results of Ophyra aenescens predation study 1 08

TABLE PAGE
15. Experimental designs of and the larval diets used in the
competition studies involving larvae of Eermetia iZlucens
vs. larvae of Musca domestica and Ophyra aenescens 115
16. Experimental design of and the larval diet used in the
competition study involving larvae of Eermetiu iZlucens and
larvae of Sarcophaga robusta 116
17. Results of the competition study between Eermetia iZlucens
and Ophyra aenescens 118
18. Results of the competition study between Eermetia iZlucens
and Musca domestica 119
19. Results of competition study between Eermetia iZlucens and
Sarcophaga 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 1 and 10 ppm 127
22. Bioassay of manure from hens fed diets containing TH 6040
and ZR-515 at 1 and 10 ppm 128
23. Data from pupal traps set in manure from hens fed diets
containing TH 6040 and ZR— 51 5 at 10 ppm 130
24. Number of pupae, number of pupae eclosed, and per cent
mortality when larvae of Musca domestica were reared in
poultry manure containing two levels of methoprene sand
granules 133
25. Number of pupae, number of pupae eclosed, and per cent
mortality when larvae of Musca domestica 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 133
28. Larval mortality and larviform pupae formation resulting
from various levels of CGA 72662 in growth media of
house flies 139
v i i

TABLE PAGE
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 ]49
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 1 60
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 1 63
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 domestica, Eevmetia illucens,
Stomcxys caicitvans, Eematcbia ivvitans, and Ophyra sp.
in two black] ight electrocutor grid traps 1 66
41. Results of knockdown tests with Farnam baits 170
42.Results of residual tests with Farnam baits
VIM

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
Malrin^M with MuscamoneTM fly baits : 1 78
46. Results of the residual test using BW 21Z and Golden
Malrin^"M with Muscamone^1'1' fly baits 180
47. Results of the attractiveness test using BW 21Z and Golden
Malrin^ with Muscamcne"^ fly baits 181
TM
48. Results of the field test using BW 21Z and Golden Malrin
with Muscamone"^ 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 1 85
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 1 88
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. LC5015 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 Ravap"^ at the
tilling site for northern fowl mite control 203
IX

TABLE PAGE
59> Formulations, mixing procedures, and application rates
for synthetic pyrethroids applied to floor birds in
Chipley, FI 207
60. Pre- and post-treatment field-estimated and converted mite
population counts and treatment means for eaoh treatment
from floor birds treated with two synthetic pyrethroids
in Chipley, FI 208
61. Daily egg production means of birds treated with two
synthetic pyrethroids in Chipley, FI 213
62. Data collection and treatment application schedule for the
Ravap"i"M northern fowl mite trial in Chipley, FI 217
63. Pre- and post-treatment mite population means by treatment
and strain from caged-layer trial at Chipley, FI 219
6A. Pretreatment mite population means by strain (treatment
ignored) and by treatment group from the caged-layer trial
at Chipley, FI 220
65. Post-treatment mite population means by strain (treatment
ignored) and by treatment group from the caged-layer trial
at Chipley, FI . . 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, FI 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 22b
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
x

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 triai at Chipley, FI 231
75* T-test of egg production treatment means by strain from
the caged-layer trial at Chipley, FI 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, FI.... 236
78. Mean operator and tractor time, and the amount of fuel
required to till one 91.4-m Ca1 ifornia-sty1e poultry
house 245

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 8l
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
x 1 1

FIGURE PAGE
19. Graphic representation of the four Ophyra aenesaens adult
longevity studies 96
20. Regression curve for data from Ophyra aenesaens predation
study 109
21. Areas on the cephaloskeleton of Ophyra aenesaens compared
with those of Musca domestica 111
22. Basal sclerite of Musca domestica 112
23. Basal sclerite of Ophyra aenesaens 112
24. Oral sclerite of Musca domestica 113
25. Oral sclerite of Ophyra aenesaens 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 1 40
29. Probit curve, fiducial limits, LC50, and regression equation
for CGA 72662 dosage-mortality data 1 43
30. Larva! 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
x i i i

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 se¿< 176
39- Probit curve, fiducial limits, and LC50 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, FI 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, FI 232
45. Plot of egg production means vs. precount mite means by
strain from caged-layer trial at Chipley, FI 237
x ( v

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, 197 S
Chairman: Dr. J. F. Butler
Major Department: Entomology and Nematology
The house fly, Musca domestica (L.), and the northern fowl mite,
Omithonyssus syluiarum (C.S 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
jm
regulator (CGA 72662 ‘ ) 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. LCsc bioassay of CGA 72662 for house fly larvae
was 0.A5 ppm.
Methoprene and dimil in were evaluated as oral larvicides, but gave
poor field results. When 20 pom 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 fly
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, Eermetia HZujcens (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. ittvjcens and M.
domestica 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 met homy 1 bait had the
longest residual and was killing at a rate higher than 25% after a
TM
o-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 1A3, BW 21Z, and SD 43775 synthetic pyrethroid compounds were
applied as a residual treatment to wooden panels hung in poultry houses,
all compounds produced 100% fly mortality 121 days post-treatment.
xv 1

In laboratory bioassay, permethrin LD50 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 Ravap 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 1 1

INTRODUCTION
In many areas of Florida, poultry farms are located in close
proximity to large housing developments. House flies (Musca domestica,
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, Cmitkonyssus sylviarum (C.S F.), can 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 jeopardy.
In this dissertation house fly control was evaluated by roto-
til ling, 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 oral larvicides. IGR's were
also evaluated when applied topically to poultry manure.
1

Qphyra aenescens (Weid.) was tested as a biocontrol agent against
house flies. Laboratory and anatomical evidence is presented to show
2
that larvae of 0. aenesoens 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, Musca domestica (L.) , is a major economic pest of
livestock and poultry. In 1977 the poultry industry in Florida lost
an estimated 55.6 million due to flies (Butler, 1973). The mere
presence of house flies in great numbers indicates the need for improved
sanitation measures (Scudder, 19^+9) and may trigger legal action.
From early Biblical times, when swarms of flies ravaged Egypt,
through ancient history (C1ouds1ey-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, 19^9). 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 dysentary, 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 (191^), 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 coth 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, 19&9b).
Bionomics
Distribution
The house fly (Musca domestica) was described by Linnaeus in 1758,
and is known as an ubiquitous insect (West, 1951)- Hewitt (191*0 called
the house fly a qualified ubiquitous insect because M. domestica 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 days

5
Some differences in the reported lengths of the life cycle can be
attributed to varience in environmental factors, such as temperature.
Melvin (193^) 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 a 1.,
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 1 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 larva! 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. domestica, 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, 1369). A female may lay up to

6
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, 19^7). 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. domestica will develop in any decaying and fermenting
organic material (James, 19^+7), in kitchen refuse and decaying vege¬
tables (Bishopp et al., 1915), and in manure of all types (Bishopp et al.,
1915; James, 19^7), 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 (Mel lor. 1919). Larvae that develop in microbial contami¬
nated media may produce adults free from contamination (Greenberg,
1973).
As in other families of díptera, males of M. domestica emerge
from their puparia before the females (Mel lor, 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., 1951). Greenberg (1973) reported a dispersion of
2.3 to 11.8 km within 24 hr.

7
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 pieces 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 Dreeding 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 wet 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 3 after about 12 hr due to

8
bacterial action (Beard and Sands, 1973). Since house fly larvae pre¬
fer a moisture level between 60 and 75% (Hiller et al., 197^) and a pH
of between 7 and 8 (Erofeeva, 19t>7) , 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., 197^), 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.

10
Spa tangía 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.
Spatangia endius, Walker (Hymenoptera: Pteroma1idae) 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 (Abies and Shepard, 197^; Legner, 1967; Legner and Brydon, 1966).
It is noted for its ability to rapidly find hosts (Abies 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 (01 ton and Legner, 1975). Be¬
sides the pupae of M. domestica, S. endius also parasitizes pupae of
Fannia femoralis and Ophyra leucostoma (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. domestica 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 muse id fly easily dis¬
tinguished from other members of the genus by its rufous-ye1iow palpi.
It was described in 1830 by Wiedemann, who placed it in the genus

Anthomyia; in 1897, Stein transferred it to Ophyra Rob ineau-Desvoidy
1830 (Johnson and Venard, 1957). Several subsequent descriptions of
the genus have been published, all placing Ophyra in the family
Anthomyiidae (Mai loch, 1923; Seguy, 1923; Aldrich, 4 928; Graham-Smith,
1916; and Bryan, 193*0. After studying the male terminalia, Crampton
(19****) 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. Ophyra 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 a I., 1965).
Biology and rearing. The biology and morphology of Ophyra
aenescens were described by Johnson and Venard in 1957- They used a
TM
larval medium consisting mainly of C.S.M.A. 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 28° 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 21° C + 1. 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).

12
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 Ophyra are predaceous. This
was supported by Kei 1in and Tate (1 930) who described the larvae of
0. leuoostoma as having buccopharyngeal armature characteristic of
larvae that are both saprophagous and carnivorous. Later experiments
proved that 0. leuoostoma was predaceous (Peck and Anderson, 1969;
Peck, 1969), but not cannibalistic (Anderson and Poorbaugh, 1964b).
Relationship with Musca domestica in poultry manure. Ophyra
leuoostoma (Wied.) occurs commonly in poultry houses in many parts of
the world (Peck and Anderson, 1970; Legner and 01 ton, 1 963; Fuji to
et a!., 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, perso-nal communication).
Hermetia illucens (L.)
Descr i pt ion . Herme tia illucens-, the black soldier fly (Sutherland,
1978), is a rather large hem isynanthiopic 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. illucens in 1738 (May, 1961). Mai loch (1917),
Ricardo (1929), and Borgmeier (1930) described the immatures and pupae.
James (1935), Linder (1938), and !ide and Mileti (1976) described the
adults, with Linder's description being the most detailed. May (1361)
described both adults and immatures.

13
Bionomics of Herirte tía. 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, I960 *
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
t i ssue (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 eclosed 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. illucens to be eurygamous. The adults reared by the former authors
did not mate, but the females laid masses of sterile eggs. Tingle et al.

(1975) succeeded in getting H. iltueens 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 Copello (1926).
Due to the variable length of the larval and pupal stages, there
are probably no more than two generations of H. iltueens 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 iltueens 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. iltueens in the Eastern
hemisphere (Barbier, 1952; Peris, 1962; Adisoemarto, 1975)- James
(I935) states that H. iltueens has been spread by commerce. Van Dyke
(1939) believes H. iltueens 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. iltueens 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. iltueens in honey bees' nests in the U.S.
Larvae have also been reported from nexts of Melponidae, a family of
stingiess bees (Borgmeier, 1930), from dead crabs (Ricardo, 1929), and
from a human cadaver (Dunn, 1916). Other habitats include beeswax,
catsup, decaying vegetables, potatoes (Mai loch, 1917), and outdoor
privies in the Southern U.S. (James, 19^+7) -

15
Myiasis. Larvae of H. illucens may cause myiasis in man, particu¬
larly intestinal myiasis due to accidental ingestion of eggs or larvae
(James, 1947; Greene, 1952; Werner, 1955).
Predators and parasites. Only one predator ofmH. 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 1952, 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 domestica (L.) in privies. The presence
of larvae of H. illucens and M. domestica 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. domestica were present
(Fletcher et al., 1956). Hypothesizing that the larvae of H. illucens
may interfere with the development of M. domestica, 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.

16
Musca domestica 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 that larvae pf M. domestica were
absent from privies where excretia was semi liquid and infestations of
H. iltucens 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 i r. soldier fly breeding.
Relationship with Musca domestica (L.) in poultry manure. The
presence of larvae of M. domestica and H. iltucens 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. iltucens and M. domestica 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.
!n the field, it was shown that H. iltucens larvae will replace
M. domestica larvae in poultry manure if the manure is moistened (Furman
et al ., 1959) . It was also demonstrated that larval populations of H.
iltucens will develop successfully when the larvae are introduced under¬
neath the crust of dry manure.

17
The outlook for H. illucens as a biological control agent in Mexico
is considered good (Vazquez-Gonza1ez et al., 1962). These authors advo¬
cate keeping poultry manure wet, especially in the dry season, and
destroying manure cones to augment H. illucens populations.
Chemica1 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.
Axtel1 and Edwards (1970) field-tested various larvicides against
larval populations of S. illucens in poultry manure. The best control
TM
was achieved wich 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. thurengiensis 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, 196A; Beard
and Wa1 ton , 1965) -

-¡8
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,
1375). 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 díptera. 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 1 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

19
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%, respect iveland had no effects
on the hens' weight. Methoprene was not effective, however, when
poultry manure was treated topically in the field.
D imi 1 i n
Dimil in, also known as TH-6040 and dif1ubenzuron, 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, 197*0. 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 (Abies 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).

20
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 acic, linseed oil, Mg2S0»,, and NaCl 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 nouse 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
(Tañada et a 1., 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
Dicnlorvos, 20% Shell Vapona resin strips ground up, gave good
control of house fly larvae and adults for about 7 weeks with three

21
treatments (Bailey et al., 1971k). Dichlorvos is also toxic to predaceous
m i tes (Axte11, 1966).
TM
Rabón , when applied to poultry manure as a larvicide, controlled
flies for 1 (Bailey et al., 1968) to 2 weeks (Matthyjsee and McClain,
1973). Rabón 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 0.2.6 g per bird in 152.0 1 of water, achieved between 68 and 94%
control of fly larvae (Jaynes and Vandepopu1iere, 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., 1371). Schreck et al. (1975) limited light trap catches to
Stomoxys aalo-itvans 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

22
increased the house fly catch 1.8- to A.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 witn 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.

23
Wicht, Jr. and Rodriguez (1970) mixed LCor concentrations of naled
JJ
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 ]%
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 a 1 . , 1 97 la).
Rogoff et al. (1964) demonstrated the presence of a house fly sex
pheromone which Carlson et al. (1971) later identified as (Z)-9~
TM
tricosene, or Huscalure . 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.

Ik
(1377) tested compounds attractive to house flies and found that
trimethylamine and indole were the main house fly attractants. Baits
consisting of trimethylamine, indole, NH^Cl , and iinoleic acid were
significantly superior to commercial preparations containing (Z)-9~
t ricosene.
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., 197*0. Permethrin was
shown to have a knockdown 8 to 16 times faster than that of allethrin,
and an LD^q 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-aerosoi system
that used a 0.7% synergized pyrethrin solution for fly control in

25
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 a!.,
1948). DDT had only recently been advocated for use on poultry farms
despite its slow knockdown and kill (Wolfenbarger and Hoffmann, 19^4).
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, methoxychlor, 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 ]% 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~fo 1d in ¡953 (LaBrecque et al., 1958), and 275*
fold in I960 (LaBrecque and Wilson, 1961).

26
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 , st. irofos, and bromophos.
TH
Ficam gave good results as a contact residual against house flies.
Sucrose was added to the solution to improve the knockdown. No resis¬
tance data are available (Lemon and Bromilow, 1977).
Synthetic pyrethroids
The first synthetic pyrethroid to be synthesized was allethrin
(Schechter et a 1., 19^9) 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. domestica than mixed esters of natural pyrethrins (Elliot et al.,
1967). Haskins et al. (197*0 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
LDj-q 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).

27
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 (Prasittlsuk and Busvine, 1977) and cattle ticks, Boophilus
microplus (Nolan et al., 1977).
Shono et al. (1978) reported that metabolic detoxification by
ester hydrolysis and hydroxy 1 at ion is a major factor limiting the
insecticidal activity of the permethrin isomers.
Northern Fowl Hites
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
communication), 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 ¡920, 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 mi1!ion annually. Smith (1978) quoted
DeVaney a; estimating an annual $66 million loss due to external

28
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 sytviarwn
(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 sylviarim (C.S 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 gatlinae. The two can be
distinguished by the shapes of the anal plates and by the shapes of the
dorsal shields (Lapage, 1956; Baker et a]., 1956; Weisbroth, 1960).
Ornithonyssus sylviariMn 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; Georg i, 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

23
0938) 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, 196k), and the»entire lire cycle
has been described (Sikes and Chamberlain, 1954; Souisby, 1968). Accord¬
ing to Sikes and Chamberlain (195*0, 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
1 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 1001. Length of the cycle varies at least partly due to the
intermittant feeding habits of the mites (Cameron, 1938).
The whole 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, 1978./,
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. sylvianan will live
from 2 to 4 weeks (Cameron, 1938; Baker et al., 1956; Kirkwood, 1363;
Loomis, 1973), as compared to 3** weeks for Dermanyssus gall-ivae
(Kirkwood, 1963).

30
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 m»ites 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. sylvianan (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 a 1. , 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. Sinc§ 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, 1953) 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 al., 1957; roulk and Matthysse, 1965; Phillis and Cromroy,
1972; Phillis et a!., 1976) ana exotic caqed birds (Sulzberger and
Kaminste in, 1936; Anonymous, 1951). Several host lists are available.
(Peters, 1933; Cameron, 193?; Strandtmann and Wharton, 1958).
Cameron (1538) lists rodents and man as accidental hosts. Other
such hosts are rabbits (Sikes and Chamberlain, 195^), the house mouse,
Mus muscuius (Drummond, 1357), the big brown bat, Evtesicus fuscas} the
cave bat, Myotis velifev (George and Strandtmann, I960), and the norway
rat, Battus noroegicus (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 Chamberlain, 195'0-

32
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-pu11et
farms, infested trucks and crates used to 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-
sem¡nation.
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 a].,
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, 1933) .
While northern fowl mites seen crawling on freshly laid chicken
eggs are an indication of a mire infestation, the number of mires

33
observed is not necessarily an indication of the severity of the infes¬
tation (J. F. Butler, 'personal oomw.n-ioo.tion). 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. A!though 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.,
197*0. 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
popu1 at ions.
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).

34
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; Metcaif et a I., 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
Al 1 ergic reactions
Gamasoi dosis, a poultry handlers1 dermatitis caused by fowl mites,
was reported in 132A (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; Ebelinq, 1975).
Riley and Johannsen (¡915/ 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 beer, used by
recent authors to describe the condition (Cahn and Shechter, 1958;

35
McGinnis, 1959; Genest, I960). Papular, ves iculo-papu1ar, 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 mite^ may not be
necessary to produce symptoms as both body parts and excretory products
of the mites have inherent toxic properties (Chandler, 19^9).
Several non-pou11ry-re 1ated cases of northern fowl mite dermatitis
have resulted from mites entering buildings via window air conditioners
(Cahn and Shechter, 1958; McGinnis, 1959; Genest, I960). 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 2k 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., 19^5, 19^6, 19^7), 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., 19^7; Hammon et al.,
19^8; Sisseru, 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
transmittor of western equine encephalitis (Chamberlain and Sikes, 1955)

36
and eliminated it as a possible transmittor of St. Louis encephalitis
(Chamberlain et al., 1957; Chamberlain, 1963).
The northern fowl mite has also been accused of transmitting fowl
pox (Brody, 1936), Newcastle virus (Hofstad, 19^9), Lankesterella oovv-i,
a blood parasite of rooks (Baker et al., 1959), a Bedsonia species of
Ornithosis virus (Meyer and Eddie, I960), and a microtatobiote, order of
Rickettsiales, of the family Bartonel1aceae (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, 1973).
Benbrook (1965) gives the most comprehensive list of controls prior to
19^0, 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 SC^ 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 SO^ should be used with
caution since it can be toxic to birds and man. Many organophosphorus

37
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 19^0, 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 2k 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.

38
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) intermittant1y
fed chickens a diet that was 5% sulfur by weight and.control 1ed 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 nothern fowl mite control
2
when added to poultry litter at the rate of 0.5 kg per 4.7 m of litter
(Foulk and Hatthysse, 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 \% sulfur spray proved ineffective
for northern fowl mite control on caged birds (Furman, 1953).
TM
Nicotine sulfate, or Black Leaf 40 , 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
interva1s.
Nicotine sulfate gave good northern fowl mite control for up to
1 month (Outright, 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

39
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 1 indane
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 (Dali et a!., 1964).
Malath ion
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, I960). Litter treatments
2
of 4% dust at a rate of 0.3 to 0.5 kg per 1.9 m 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 concentrar, ions of malathion.
A 4% solution killed all birds dipped including one mature goose (Furman
and Weinmann, 1956).
Ca rba ry1
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, I960). 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).
Ronne1
Laboratory and field tests have shown that ronne! is more toxic to
northern fowl mites than either barthrin (a botanical) or malathion

41
(Bigley et al., I960). 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, I960; Foulk and Matthysse, 1963).*
Mi seellaneous compounds
Other compounds, mostly organophosphorous compounds, giving good
northern fowl mite control are coumaphos (Bay 21/199) sprays and dusts
(Kraemer, 1959; Hoffman, I960; Knapp and Krause, I960; 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 (Dylox"^)
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 pyrethroids,
TM
Ectiban 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

42
or practical value of either one is questionable. Volck, which is
commonly used on plant pests and is nontcxic to birds and mammals, was
tested as an ectoparasite control agent for farm animals (Bruce, '928;
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).
System;cs
Su 1faquinoxa1ine alone or with other sulfonamides acts as a systemic
acaricide in birds infested with northern fowl mites (Beesley, 1973).
When feed containing 0.033% su 1faquinoxa1ine was fed to layers, mite
populations were reduced to near zero in approximately 4 weeks (Furman
and Stratton, 1963). Feed with 0.05% sulfaquinoxa1ine 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 su1faquinoxa1ine,
sulfadimedine, su 1 famerezine, and su1fathiazo1e 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 AMD MATERIALS
Laboratory Trials
Environmentally Controlled Rearing Conditions
TM
Rearing of immature diptera was accomplished in a Percival
forced-air, upright growth chamber. The temperature was maintained
at 29.4 C, and continuous lighting was provided by two AO-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.^ C, continuous iighting,
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 wi11 be referred to specifically throughout
the text ¡f applicable.
Colonization and Rearing of Flies
Musoa domestica (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 CSMA' (Consumer Specialties Manufac¬
turing Association) and allowed to oviposit. Jars were screened and
43

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 (¡928) and Monroe (1962).

45
Ophyra aenescens (Wi ed.) -- 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
mea 1 .
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.
Hermetic illucens (L.) — the black soldier fly. This fly was
reared in the laboratory on many occasions, but attempts to colonize it
did net 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,
E. illucens was reared in chicken feed, fish meal, and mixtures of fish
meal and CSMA, all moistened with water.
Phormia regina (Me i gen) -- 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

46
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 Ophyva aenesaens.
Fannia oan'tcvlaris (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.
Saraophaga 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 l80 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

47
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 (1947), revealed that all
specimens belonged to the same genus and species, Saroophaga robusta
(Aldrich), syn. S. plinthopyga (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 Cepha1oske1etons of Third-Instar Fly Larvae
Cepha1oske1etons 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 cepha1oske1etons had dissolved. At the com¬
pletion of the KOH treatment, larvae were dried for 1-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 cepha1oskeletons were stored overnight in phenol and
the remaining portions of the larvae were discarded.
Cephaloskeletons were worked into a mixture of phenol-ba1sam and
placed in desired positions on mounting slides. Care was taken that
the specimens were completely covered with the phenol-ba1sam mixture.

48
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 tc 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
covers!ips are added before phenol has completely evaporated, specimens
may be damaged.
After the phenol had evaporated, covers!ips 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-ml 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 (1GR) 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 1GR.
Control diets were prepared using plain water.
Diets were placed in 360-ml 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 eclosed 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.

50
Attractiveness tests. Baits were sprinkled into bait stations
fashioned of 3.8-1 milk jugs (R. C. Axtel1, 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 2k 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, ail 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 2k hr.
Laboratory colony house flies, 3 to 5 days old, were immobilized
with a vacuum and males discarded. While immobilized, female Hies
TM
were dosed with test concentrations using a 10-yl Hamilton syringe
TM
equipped with a Hamilton 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 2k hr. Criterion for death was total cessation of
movement.

51
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 viere treated with 100 ml of acetone. Next, pipets were tapped
on paoer towelling for 2 min, tapered ends down, to dry the insides.
TM
More complete drying was achieved by using a Gast electric pump to
force air through the pipets for 20 min. Pipets were removed from the
pump and used within 1 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

52
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
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

53
Figure 1. View of tractor and tiller

54
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 Ca1 i fornia-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 Ca1 ifornia-sty1e houses were
used in the pest management studies. The layout of the farm and the
designation of the houses a re shown in Figure 3- The watering system
TM
consisted of one Hart 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 3! 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.

55
Figure 2. Tiller in operation.

Egg cooler
Figure 3. Layout and numeric designation of pouI try houses at the tilling site.

57
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 wee 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

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

59
of five 360-ml 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

60
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-davs. 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

61
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 l). The basal diet was mixed in
TM
136.4-kg lots in a Kelley-Duplex vertical mixer. When the basal diet
was thoroughly mixed, ca. A.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 ali
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.

62
Table 1. Composition of basal diet for poultry feed trials.
Ingredients
Per cent
Yellow corn
69-33
Soybean meal (50% protein)
19.00
Alfalfa meal (20% protein)
2.50
Ground 1imestone
6.17
Dicalcium phosphate
2.25
Iodized salt
.25
Micro-ingredient mix3
.50
Tota 1
100.00
C.M.E./kgk
2890
c
Per cent procein
16.2
c
Per cent calcium
2.98
Per cent phosphorus0
.73
Supplies per kg of diet; 6600 I.U. vitamin A; 2200 l.C.U. vitamin
D,; 11 I.U. vitamin E; 2.2 mg menadione dimethy1pyrimid i na 1 bisulfite
(flPB); k.k mg riboflavin; 13-2 mg pantothenic acid; 59-6 mg niacin;
998.8 mg choline chloride; 22 meg vitamin 3^5 110 meg biotin; 125 mg
ethoxyquin; 60 mg manganese; 50 mg iron, 6 mg copper; 0.198 mg cobalt;
1.1 mg iodine; 60 mg zinc.
b Calories metabolizable energy per kilogram.
Q
Calculated values.

63
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-ml glass jar. Amounts were
preweighed and applied to manure treatment blocks until the shaker was
empty.
Mixing and Application of Liquid 1GR1s and Organophosphorus Larvicides
Liquid !GR1s and commercial larvicides were applied to manure with
TM
a 7.7-1 Sears 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-ml 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 1GR 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 1GR 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

64
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 i0% 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.

65
Figure 5* Light trap opposite egg processing
Note the fiat-deck cage arrangement
room.

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

67
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).

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

69
Compounds were mixed using formulas as described in Neal (197*0
and applied to run-off with hand-held triggei—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 k 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 mi tes 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

70
Figure 8. A pair of workers examining a hen for mites.
The counter is on the left. Note the
stair-step cage arrangement.
Figure 9- A close-up view of Figure 8. The darkened
area on the chicken is due to mites and
mi te debris.

71
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.
Est¡mat ion
Estimation
Field Est.
100
2710
27.10
500
2895
5.79
1000
5065
5.07
2500
4735
1.90
5000
4050
0.81
x|
II
CO
CO
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 8.S G. cans and a
Sears 7-7-1 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.

72
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.S 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, 197*0. 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 ¡n-depth explanation of
probit analysis and the calculation of a probit line was found in
Finney (196*4). 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).

Table 2. Compounds utilized for fly and/or mite control.
Common
Code letters and
Chemica1
name
numbers and/or trademarks
name
Cooperator
Insect Growth Regulators (IGR's)
methoprene
ZR-515, Altosidâ„¢
¡sopropy 1 (E , E)- 11-met hoxy-3,7,11-
trimethy 1-2,4-dodecadienoate
, TM
Zoecon
dimi 1 i n
TH 6040
1 -(4-ch1oropheny1)—3~(2,6—
dif1uorobenzoy1) urea
Thompson-Hayward
CGA 72662
La rvicides
Ciba-Geig y
dimethoate
r TM
Cygon
0,0-dimethyl S-(N-
methy1 carbamoyl methyl)
phosphorodithioate
—
dichlorvos
DDVP, Vapona^'
2,2-d i ch1oroviny1 dimethyl
phosphate
*
d â„¢
Ravap
TM
Rabón [2-chloro-1 -(2,4,5
trich1oropheny1) vinyl dimethyl
phosphate] + Vapona^M (See
dichlorvos above)

Table 2.
Con t¡nued.
Common Code letters and Chemical
name numbers and/or trademark name Cooperators
Baits
—
TM
Bomy 1
dimethyl 3-hydroxyg1utaconate
dimethyl phosphate
Farnam
d1ch1orvos
See Larvlcldes above
ronne1
TM
Kor1 an
0,0-d¡methyl 0-2,4,5“tri chioropheny1
phosphorothioate
Fa rnam
methorny 1
. „ TM
Lannate
methyl N-[(methy1carbamoy1)oxy]
thioacetimi date
Farnam
Golden Ma1r1nâ„¢
dichlorvos (See Larvicides above) +
ronne1 (Above)
Contact Residuals
fenva1erate
SD 43775
—
Shell
permethrin
BW 21 Z
(3*phenoxypheny1)methy1(T)-cis,
trans-3~(2,2-d¡chioroetheny1)-2,2-
dimethyl eye 1 opropanecarboxy1 ate
Burroughs Wellcome
permethrin
ICI 143, JFU 5819,
JFU 5021A
—
ICI Americas
''j
-p-

Table 2. Continued.
Common
name
Code letters and
numbers and/or trademark
Chemica1
name
Cooperators
Acaricides
fenvalerate
SD *13775
—
She 1 1
resmethrin
SBP-I382
—
Penick
permethrin
BW 21 Z
(See Contact Residuals above)
Burroughs Wellcome
—
Ravap
(See Larvicides above)
—
permethrin
TH
Ectiban
(3-phenoxypheny1)methy1(t)-cis,
trans-3_(2,2-dichloroethenyl)-2,2-
d¡methy 1 eye 1opropanecarboxy 1 ate
1 C 1 Uni ted States
carbary1
c . TH
Sev 1 n
1-naphthyl methyl carbamate
—
mal athion
TH
Cyth1 on
diethyl mercaptosuccinate, S-ester
with 0,0-dimethyl phosphorodithioate
fe-—

RESULTS
House F]i es
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 everv house twice a day, 7 days a week. Houses 1 through A
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.
ResuIts. 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, ,i f 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.
76

77
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.

78
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.

8o
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 1evel.
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.
Resu1ts. 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

8i
A
B
Figure 12.
1 2
3 A
pq
4
'Va-
B
C
c
—
B
0
S
N
B
T
C
R
B
0
S
L
B
lLJ
C=CHIPS
S=SAND
B=BLANK
Experimental design for adding builder's sand and wood
chips to houses 1 through k at the tilling site.
Only the front two-thirds of the houses are shown.

82
Figure 13. The appearance of chips before spreading.
Figure 14. The appearance of chips after the initial
tilling.

83
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

84
ca. 4 cm below the lower surface of the concrete walks. Subsequent
heavy rains flooded the farm and the manure collection areas (Figure
15)* It was decided to add wood chips to all manure collection areas
to help dry and texturize the manure. The treatment schedule is shown
below:
House Chips
1 not added
2 added to rear of house only
3 added to entire house
4 not added
The level of the chips before tilling is shown in Figure 16. After
chips were added, all houses were tilled twice to stir in the chips and
make moisture levels more uniform when the pretreatment manure samples
were taken. House 1 was tilled on Mondays, house 3 on Mondays and
Fridays, and house 4 on Mondays, Wednesdays, and Fridays. House 2 was
not tilled.
On Fridays, one 360-ml manure sample was collected from the center
of each quarter of each house. At the laboratory, 30 g of each sample
were weighed on an electric pan balance and dried for 36 hr on Berlese
funnels. Samples were reweighed and the dry weights subtracted from
30 g. These differences were converted to per cents to arrive at the
moisture contents of the samples.
Resu1ts. The results from the manure samples dried for moisture
content are shown in Table 3 and Figure 17.
Moisture levels were reduced in all treatments except 2-B, where
no chips were added to the manure and the manure was not tilled. The
moisture level in this treatment increased significantly from 49*35 to
64.32% and constituted a net moisture increase of 14.97%. All other
changes in moisture levels were nonsignificant.

85
Figure 15. The appearance of manure collection areas
at the tilling site after manure removal
and subsequent flooding. Note that the
walk has been exposed from improper manure
removal procedures.

86
Figure 16. Addition of wood chips to flooded manure
col lection areas.

87
Table 3- Moisture 1evels(%) of manure samples from the tilling site
and net change in moisture content(%).
Pou1 try
Hou ses
T reatment
1
2
3
4
Period
A
B
A
B
A
B
A
B
Pre-
Treatment
64.70
67-93
63.57
49.35a
53.30
63.10
72.83
65.13
1
68.40
57-47
67.80
64.37
66.67
61 .03
70.07
66.30
2
69-47
62.50
63-93
66.03
43.33
51.27
66.47
69.77
3
63.20
66.20
62.77
61.40
57.37
52.07
65-75
70.00
4
54.32
63.90
57.85
48.25
53.08
61.65
69.03
66.98
5
55.83
60.67
61.33
50.92
66.57
64.42
71 .57
63 25
6
59.13
61.37
58.85
53.62
54.23
59.68
63-93
67.85
7
58.50
55.55
64.47
49.02
54.93
53.80
69.63
66.78
8
47.38
51.85
44.82
55.42
41.33
49.63
55.27
46.32
9
55.20
62.58
56.45
64.32a
44.70
57.37
61.82
60.72
Net Change
-9-50
-5.35
-7.12
+14.97
-8.60
-5.73
-11.01
-4.41
Indicates significance at p < .05% between the pretreatment
sample and the sample from period 9.
Note: The duration of each treatment period was 7 days. Treatments
are as follows: 1A - no chips, tilled lX/wk
IB - no chips, tilled IX/wk
2A - chips added, no tilling
2B - no chips, no tilling
3A - chips added, tilled 2X/wk
3B - chips added, tilled 2X/2k
4A - no chips, tilled 3X/wk
4B - no chips, tilled 3X/wk

lo ISTURE (%)
80
70
60
50
40
PRETREATMENT TREATMENT PERIOD 9
2X/wk
Trt
Chips
Tilling
% A
o
- 2B
No
No
+1^.97
O
- IB
No
lX/wk
-5.35
©
- ¿tA
No
3X/wk
-11.01
A
- ^B
No
3X/wk
-4.41
â–¡
- 3B
Yes
2X/wk
-5.73
©
- 2A
Yes
No
-7.12
o
- 1A
No
lX/wk
-9.50
-8.60
I
Treatment Period
Figure 17. Net results of the manure drying experiment with wood chips and tilling 9 weeks post-
t rea tment.
CO
co

39
The greatest reduction in moisture level, 11.01?, was in treatment
block 4-A, where no chips were added and manure was tilled 3 times per
week. This treatment area also had the highest moisture level when the
experiment began and third highest when the experiment ended. The final
moisture level of treatment block 4-A was approaching 60? which is the
minimum level that flies prefer to use for breeding purposes (Miller
et a 1., 1974).
Moisture levels of treatments 1-B (without chips), 4-B (without
chips), 1-A (without chips), and 3-B (with chips) were all reduced to
near 60? or below 60? by the use of different tilling schedules (Figure
17). The moisture level of treatment 2-A (with chips) dropped to below
the 60? level during the treatment period with no tilling at all. The
moisture level of 53.30? in treatment 3”A (with chips) was well below
the 60? level when the experiment began and dropped to 44.70? when the
experiment ended. Figure 18 shows manure in 3"B at the end of the 3~
week experiment.
When the experiment was terminated, the manure in all treatment
areas except 4-A, 4-B, and 1-B maintained good form after tilling. The
manure in the three above-mentioned areas appeared quite wet even though
the moisture levels were just above 60?.
In treatment areas that were not tilled, a crust had formed on the
manure in most places. In wet areas, this kept moisture in and pre¬
vented further drying. Pockets of house fly larvae were not noted in
these areas during the course of the experiment.
Arriving at more accurate manure moisture levels was limited by the
number of samples that could be processed by our laboratory. Only two
samples could be taken from each treatment area every week. Eight

Figure 18. The manure in 3~B at the end of the
experiment.

31
samples would have been much more satisfactory since each treatment area
was 45.7 m in length. Overall limiting factors were drying facilities,
driving distance to the tilling site, and labor.
Ophyra aenescens Basic Biology Studies
Larval diet and adult longevity studies. Four preliminary studies
were used to develop larval and adult diets for 0. aenescens. The
ingredients in the two larval diets are shown in Table 4, and the diet
combinations for larvae and adults are summarized in Table 5. Since
these were only preliminary studies and the resulting fly numbers were
relatively large (> 30), there were no treatment replications.
Pans containing the experimental larval diets were seeded with 500
and 1000 eggs. Eggs were covered loosely with approximately 2 to 3 cm
of medium to simulate oviposition. Pans were covered with fine nylon
screen and placed in the growth chamber (29-4 C) .
After 10 days, pupae were separated from the media by flotation,
dried, and placed in standard colony cages. Each cage contained one of
the adult diets shown in Table 5, and a water source with polyfoam chips
on the surface to reduce drowning mortality. Cages of adults were kept
in the walk-in growth chamber (26.7 C).
Adults were allowed to emerge for 24 hr, after which the remaining
pupae were removed from the cages. Beginning 24 hr later, dead flies
were removed from the cages on a daily basis and daily mortality records
were kept by sex until all the flies in each cage were dead.
Results. The results of the four 0. aenescens adult longevity
studies plus the average adult life span and length of the life cycle
in each study are shown in Table 6. Raw data are shown in Appendix 1.
Fortification of the larval diet did not shorten the 12- to 14-day

92
Table 4. Fortified and unfortified diets used during the preliminary
colonization studies with Ophyra aenescens larvae.
Constituent
D i et
Unfort ified
Fort ified
CSMAâ„¢3
500 ml (151g)
400 ml (113g)
Horn fly dry mix^
—
100 ml (72g)
Water
375 ml
275 ml
Yield
~700 ml
-800 ml
Consumer's Specialties Manufacturing Association (Ra1ston-Purina).
k Horn fly dry mix (Greer, 1975) "wheat flour~960g, fish meal-720g,
Na2C03-i20g, alfalfa meal-400g.

93
Table 5- Combinations of larval and adult diets used during the four
preliminary colonization studies with Ophyra aenesaens.
D i et
1
2
3
4
Larva 1
Unfort ifieda
Fortified^
Unfort rfied
Fortified
Adu 11
Nasogastric mix
and fish meal
Nasogastric mix,
fish meal, and
cane sugar
Nasogastric mix,
fish mea 1, and
cane sugar
Fish mea 1
and cane
suga r
3 T M
CSMA ' (Ra1ston-Purina) plus water.
k CSMA1^, horn fly dry mix (Greer, 1975)— 96Og wheat flour, 720g
fish meal, 120g Na2C03. 400g alfalfa meal; plus water.
C Challenge Cream and Butter Ass., Los Angeles, Ca. Each 270g
furnishes: 7*5g moisture, 2.7g milk fat, 73-Og protein, 16.6g ash,
168.Sg carbohydrate, 1.Og sodium, 3-2g potassium, Img thiamine hydro¬
chloride, 2mg riboflavin, 1Omg nicotinamide, 0.5mg pyridoxine hydro¬
chloride, 1Omg calcium pantothenate, 0.25mg folic acid, 150mg ascorbic
acid.

Table 6. Summary
and the
of the four Ophyra
1ength of the 1 ife
aenesoens adult longevity
cycle from each study.
studies inc1ud i ngaverage adult 1
i fe span
Diet
1
2
3
4
Larva 1
Unfort ifieda
. c . . b
Fortified
Unfort ified
Fort ified
Adult
c
Nasogastric mix
and fish mea 1
Nasogastric mix,
fish mea 1, and
cane sugar
Nasogastric mix,
fish mea 1 , and
cane sugar
fish mea 1
cane sugar
and
Hales Females
Males Females
Males Females
Males Females
Average adult
1ife span(days)
2 2
18 21
18 22
14
17
Time from egg
to adu11 (days)
13
14
12
12
3 "FM
CSMA (Ralston-Pur¡na) plus water
/
h TM
CSMA , horn fly dry mix (Greer, 1975)—960g wheat flour, 720g fish meal, 120g Na2C03, 400g
alfalfa meal; plus water.
c
Challenge Cream and Butter Ass., Los Angeles, Ca. Each 270g furnishes: 7.5g moisture, 2.7g
milk fat, 73-Og protein, l6.6g ash, l68.8g carbohydrate, 1.Og sodium, 3-2g potassium, 0.5mg pyridoxine
hydrochloride, 1Omg calcium pantothenate, 0.25mg folic acid, 150mg ascorbic acid, 1mg thiamine hydro¬
chloride, 2mg riboflavin, 1Omg nicotinamide.

95
period required for larval development, but sugar increased the average
adult life span 9 times. The average life span for adults when sugar
was not included in the diet was 2 days compared to 18 days for adults
having access to sugar. By the time adults were 2 weeks old, mortality
averaged 20% in groups consuming sugar and 90% in the group not consuming
sugar (Figure 19).
Females lived for an average of 20 days and outlived males by an
average of 3 to 4 days. Individual males and females were kept alive
in the laboratory for 36 and 39 days respectively. Sex ratios were
50:50 when the totals of both sexes from all four tests were combined
and subjected to a chi square analysis.
Larval viability study. To evaluate the effects of fortified and
unfortified larval diets on larval development, 700 ml of each diet were
loosely placed in four 1000-ml pyrex beakers. Sixty first-instar larvae
of 0. aenescens were added to each beaker. Beakers were covered with
fine nylon screen and placed in the growth chamber. When adults began
to emerge, remaining pupae were separated from the media by flotation,
dried, weighed, and set aside for emergence of remaining adults. After
adults emerged, empty pupal cases were reweighed.
Resu1ts. The number of pupae, the per cent pupation, the numerical
and per cent larval emergence, and the larval viability of 0. aenesaens
reared in fortified and unfortified larval diets are shown in Table 7.
Fortification of the larval diet significantly increased the number of
pupae that formed and the number of adults that emerged. The per cent
viabilities of the larvae grown in the fortified and unfortified diets
were 52.5 and 3-8% respectively. Development time from larvae to adults
was 14 days for both diets. There was no significant difference in pupal

4 8 12 16 20 24 28 32 36 40 44 48
Day
Figure 13. Graphic representation of the four Ophyra aenesoens adult longevity studies.
VD
CTv

97
Tabl i
e 7
The number of pupae, the per cent pupation, the numerical and
per cent emergence, and the larval viability of Ophyra
asnescens reared in fortified and unfortified larval diets.
Number of
°/
'O
Numerica1
°/
'0
Larval
Rep
Pupae
Pupation
Emergence
Emergence
V iabi1ity[%)3
Fort ified Diet^
1
41/60°
63.3
33/41d
92.7
63-3
2
34/60
56.1
30/34
88.2
60.0
3
33/60
55.0
31/33
93.9
51.7
4
29/60
48.3
27/29
93.1
45.0
137/240
57.1
126/137
92.0
52.5
Unfortified Diet
e
1
2/60
3.3
1/2
50.0
1.7
2
5/60
8.3
3/5
60.0
5.0
3
3/60
5.0
3/3
100.0
5.0
4
4/60
6.7
2/4
50.0
3-3
14/240
5.8
9/14
65.7
3.8
a
_ , number
of adu1ts
Equa1s r
number
of pupae
X 100
b
TM
CSMA , horn f
1y dry mix
(Greer, 1975)— 960g wheat
flour, 720g
fish
meal, 120g Na2CC>3 , 400g alfalfa meal;
plus water.
c
Represents the
number of
pupae that
formed from 60
f i rst-instar
0. asnesaens larvae.
d
Represents the
number of
adults that
eclosed from
the respective
number
of pupae shown
in column
2.
e
CSMA^"'1 (Ra 1 ston-Pur i na)
plus water.

38
weights due to the diet (Table 8), although pupae from the unfortified
diet were numerically lighter by an average of 0.3 mg. This indicated
that adults from both treatments were the same size.
Larval density study. This study was designed to determine whether
a minimum number of 0. aenesaens larvae must be present per unit area in
order to have maximum development in a minimum amount of time. The
fortified diet (Table 4) was placed loosely in 360-ml plastic cups to
within 5 cm of the rim. Four population levels were used, each repli¬
cated four times. One cup with media served as a replication. To each
cup in the respective treatment group, 5, 25, 50, and 100 first-instar
0. aenesaens larvae were added. Larvae were 12 to 18 hr old with an
average length of 1.0 mm. After addition of larvae, cups were covered
with screen and placed in the growth chamber. Pupal cases were separated
from the medium after the emergence and death of the adults.
ResuIts. The number of adults that emerged when the larval density
averaged five larvae per 360 ml of medium was significantly lower than
the number of adults that emerged from treatments with higher larval
densities (Table 9)* The treatment with the lowest larval density
averaged 40.0% viability while treatments with 25, 50, and 100 larvae
per 360 ml of medium averaged 91-0, 89-5, and 86.3% respectively. Results
indicate the existance of an 0. ae?iescens minimum population density
threshold in between 5 and 25 larvae per 360 ml of medium. Additional
research is needed to more accurately define the threshold.
Temperature of larval medium. Two aluminum pans, 25 cm in diameter
and 8 cm deep, were filled to within 2 cm of the rims with the forti¬
fied larval diet (Table 4). One pan was seeded with ca. 500 eggs of
0. aenesaens and the other was used as a control. Pans were screen-

99
Table 8. Uneclosed pupal weights of Ophyra aenesaens reared in fortified
and unfortified diets.
Fortified Diet3
Unfortified Diet*3
RepsC
‘Reps
d
1
2
3
4
1
2
3
4
0.0114s
0.0100
0.0116
0.0156
0.0096
0.0130
0.0119
0.0134
0.0127
0.0135
0.0114
0.0140
0.0148
0.0121
0.0106
0.0128
0.0134
0.0123
0.0128
0.0111
0.0110
0.0131
0.0135
0.0102
0.0110
0.0105
0.0111
0.0106
0.0146
0.0098
0.0131
0.0133
0.0126
0.0123
0.0123
0.0115
0.0136
0.0123
0.0122
0.0130
0.0143
0.0123
0.0125
0.0125
0.0144
0.0100
0.0111
0.0175
0.0125
0.0117
0.0144
0.0127
0.1297
0.1152
0.1219
0.1323
0.0244
0.0361
0.0356
0.0503
x =
0.0125
x =
0.0122
3 CSMA'M (Ra1ston-Purina), horn fly dry mix (Greer, 1975)— 960g
wheat flour, 720g fish meal, 120g Na2C03, 400g alfalfa meal; plus water.
° CSMA™ plus water
C Ten pupae from each beaker were chosen at random for weighing.
^ All available pupae from each beaker were weighed.
6
All pupal weights are in g.

100
Table 9- Emergence of adults of Ophyra aenesoens when various numbers of
first-instar larvae were reared in the same volume of growth
mediurn.
Rep
5
Number of first-
25
instar larvae/rep
50.
100
d1 ?
o' ?
?
d* ?
1
0 0
14 11
22 24
41 38
2
1 0
11 9
23 19
48 48
3
2 2
10 12
20 25
39 46
4
_L JL
18 _6
18 28
n 5±
E by sex
4 4
53 38
83 96
162 183
£ by trt
8
91
179
345
X
2.0a
22.8
44.8
86.3
% viability
40.0
91.0
89.5
86.3
Indicates that this value is significantly different from others
in the same row at p < .05%.

101
covered and placed in the growth chamber. After 2b hr, the temperature
TM
of the medium in each pan was measured by use of a Taylor dial thermo¬
meter. Pans were divided into quarters and temperature measurements
taken from the centers of the pans and from the centers of the quarters.
The temperatures of the media were recorded every 2b hr from day 1, i.e.
2b hr after eggs were set, through day 7 of the larval development
period. On day 7, pupae in the treated pan were floated and the experi¬
ment was terminated.
Results. The daily temperatures of larval media as influenced by
0. aenescens larvae over the 7-day larval development period are shown
in Table 10. The medium with larvae showed a daily increase in tempera¬
ture, reaching b2.b C on day 2 and then declining to 30.0 C on day 7.
The medium without the larvae also peaked on day 2 at 33-7 C and declined
to 28.7 C on day 7- Elevated temperatures in the medium with the larvae
were attributed to larval interaction with the medium. Daily differences
in temperature between the two pans were all significant (p < .01).
Competition study between Qphyra aenescens and Afusca domestica. In
this trial, 180-ml cups containing 90 ml of diet were used with four
replications per treatment. First-instar larvae of both fly species
were used to seed the diets. Experimental design and diet used are
shown in Table 11. Treatments consisting of only one fly species were
used as controls. Cups with larvae were covered with screen and placed
in the growth chamber. Adults were allowed to eclose and die before
they were counted.
ResuIts. The results of the competition study between 0. aenescens
and M. domest-ica are shown in Table 12. In all treatments where house
flies and dump flies were developing together, house fiv mortality was

102
Table 10. Daily temperatures of larval media as influenced by Ophyra
aenesaens larvae over the 7-day larval development period.
Rep
1
2
3
Day
4
5
6
7
Mediurn
with Larvae
1
45.6a
45.0
41.1
42.2
32.2
32.2
30.0
2
43.3
40.0
41 .1
34.4
32.2
31.1
30.0
3
40.0
40.0
34.4
34.4
34.4
32.2
30.0
4
40.0
42.2
38.9
36.7
34.4
32.2
30.0
5
42.2
45.0
40.0
37.8
31.1
31.1
30.0
X
42.2A
42.4A
39- 1A
37.1A
32.9A
<
OO
ro
30.0A
Med i urn
wi thout
La rvae
1
33-3
31 .7
30.0
32.2
28.9
27.8
30.0
2
28.3
35.6
32.2
28.9
31.1
30.0
28.9
3
30.0
33.3
31.1
26.7
28.9
27.8
28.9
4
32.2
34.4
30.0
26.7
27.8
27.8
27.8
5
31.1
33.3
31.1
27.8
28.9
27.8
27.8
X
31 .IB
33.7B
30.1B
28.5B
29.1B
28.2B
28.7B
All values are in degrees centigrade.
Note: Means in the same column with uncommon letters are signi¬
ficantly different (p < .01).

103
Table 11. Experimental design of and the larval diet used in the compe¬
tition study Involving larvae of Ophyra aenescens and larvae
of Musca domestica.
Treatment No.
T reatment
1
15a
0.
i b
aenescens larvae
2
15
M.
domes tica larvae
3
15
0.
aenescens larvae + 15 M. domestica
larvae
4
15
0.
aenescens larvae + 30 M. domestica
1arvae
5
30
M.
domestica larvae
6
15
0.
aenescens larvae + 45 M. domestica
larvae
7
45
M.
domestica larvae
Larva 1
TM
Diet: 400 ml CSMA
100 ml horn fly dry mixC
275 ml water
ca. 700 ml medium
a Represents the number of larvae per replication with four
replications per treatment.
b
First-lnstar larvae of both species were used in the trial.
960g wheat flour, 720g fish meal, 120g Na2C03, 400g alfalfa meal
(Greer, 1975).

Table 12. Results of the competition study between Ophyra aenescens and Musca domestica.
Reps
1
2
3
Treatments
<1
5
6
7
15 0. a
15 M. b
15 0.
+ 1 5 M.
15 0.
+ 30 M.
30 M.
15 0.
+ *t5 M.
'i5 M.
aenesoens
domestica
aenesoens
domestica
aenesoens
domestica
domestica
aenesoens
domestica
domestica
i
15
15
15
0
15
0
30
15
it
1|2
2
15
15
15
0
13
0
30
15
3
it5
3
15
15
15
0
15
0
28
15
5
^3
'i
1 if
1 15
0
15
2
30
15
5
X
|it.8c
l 15.0
o.ox
1 â–  5
0.5V
29.5Y
15.0
4.3Z
1i3.8Z
% Horta1
ity 1.7
1.7
0.0
100.0
3.3
98.3
1.7
0.0
90.6
2.8
Indicates number of 0. aenescens larvae used in each rep.
'J Indicates number of M. domestica larvae used in each rep.
Q
Represents the mean number of viable individuals at the end of the experiment.
Note: Pairs of means having common letters are significantly different (p < .05).

105
significantly higher than in the control groups, while dump fly mor¬
tality remained unaffected. Fifteen 0. aenescens larvae prevented the
development of an equal number of M. domestica larvae, and produced
mortality rates of 98.3 and 90.6% when reared with 30 and 45 house fly
larvae respectively. House fly larvae that pupated in treatments 3, 4,
and 6 reached the adult stage (Table 12). The remaining house fly larvae
viere prevented from reaching the pupal stage and died as larvae, but no
dead larvae could be found in the growth media. This strongly suggested
that the larvae of 0. aenescens were preying upon the larvae of M.
domestica.
Predation study with Qpkyra aenescens. In this experiment, adapted
after Peck (1969), the competition studies described above were modified
and the availability of the larval medium as a food source was removed.
Replications were 180—m1 clear plastic cups containing 90 ml of vermicu-
lite. Each treatment was replicated four times. Prior to addition of
larvae, enough water was added to the vermiculite to make it damp but
not soggy. Too much water forced larvae to the surface, not enough
water caused them to stick to the sides of the cups. Water was added to
all cups daily throughout the experiment to maintain the proper moisture
level. The experimental design is shown in Table 13- In treatments i,
2, and 3, 10 first-ins tar larvae of 0. aenescens were placed in each
cup of vermiculite. Then 25, 100, and 200 first-instar house fly larvae
were added daily to each cup in the respective treatment group until
pupae were noted in the cups. Treatments 4 through 8 were used as
controls. Cups were covered with screen and kept in the growth chamber
except when larvae were being added. At the end of the experiment,
flies were allowed to eel ose and die before they were counted.

106
Table 13- Experimental design of Ophyra ctenescens predation study.
Treatment No.
T reatment
Larvae added initially Larvae added daily
1
10a
0.
b
aenescens
*25a
M.
domestica
2
10
0.
ctenescens
100
M.
domestica
3
10
0.
aenescens
200
M.
domestica
4
25
M.
domestica
—
5
100
M.
domestica
—
6
200
M.
domestica
—
7
100
0.
aenescens
—
Represents the number of larvae per replication with four
replications per treatment.
b
Larvae used were first instars of both fly species.

107
Resu1ts. The results of the predation study are shown in Table 14.
The mortality rate of 0. aenesoens larvae was reduced significantly in
all cases by the daily addition of house fly larvae. Control mortalities
were 100% in all cases, indicating that neither fly species is canni¬
balistic and that in order to survive, larvae of 0. aenesoens were
preying upon larvae of M. domestica. This was, in fact, the case and
it was observed on numerous occasions.
Ophyra larvae never fed for extended periods on a single house fly
larva. After killing and feeding briefly on one, they moved on in search
of another. When a house fly larva was killed by Ophyra, other house fly
larvae were attracted to the site and began feeding on the dead larva.
The clustering of house fly larvae aided Ophyra larvae in their search
for prey. This hypothesis was corroborated by fitting the data in
Table 14 to a binomial regression model and plotting the graph in Figure
20. The goodness of fit was highly significant (p < 0.0001). Larvae of
0. aenesoens can destroy more than 20 first-instar house fly larvae per
day during the larval developmental period, but further testing is
necessary to arrive at a more accurate number. On at least one occasion,
a third-instar larva of 0. aenesoens was seen feeding on a pupa of M.
domestica that had not completed the tanning process.
Since Ophyra adults were allowed to emerge prior to processing, no
pupal weights were available to check for differences in the sizes of
the adults. An attempt was made to find a correlation between eclosed
pupal weight and adult weight, but results were negative. Adults
raised in vermiculite did not appear to be smaller than adults from
larvae raised in the usual manner.

Table Wi. Results of Ophyra aenesoens predation studies.
Treatments
Reps 1
2 3
5
5
6
7
8
10 0. + 25 M.
aeneaaerut domestica
h 10 0. + 100 M. ( 10 0. 0+ 200 M.
aenesoens domestica aenesoens domestica
10 0.
b a
aenesoens
2 5 M.
domes ticac
loo n.
domestica
200 M.
doma Li i, i- qcl
100 0.
a
aenescenc
i
1 0
6
0 9 0
0
0
0
0
0
2
2 0
6
0 9 0
0
0
0
0
0
3
3 0
5
o 8 0
0
0
0
0
0
'i
1 0
8
070
0
0
0
0
0
X
i.8Ad 0.0
6.3A
0.0 8.3A 0.0
0.0A
0.0
0.0
0.0
0.0
% Mortal 1ty 82.5 100.0
37-5
100.0 17.5 100.0
100.0
100.0
100.0
100.0
100.0
a
Indícales the number of 0.
aenencfínu
larvae initially placed in each rep
of treatments
1, 2, 3, <1,
and 8.
b
Represents the number of M.
. domestica
larvae added daily to each rep of
treatments 1 , 2
, and 3.
c
d
Indicates the number of M.
domestica
larvae initially placed in each rep
of treatments
5, 6, and 7
Represents the mean number of viable Individuals at the end of the experiment.
Note: Means having common letters are significantly different (p < .05).

Mean Number of Ophyra Adults Eclosed
Figure 20. Regression curve for data from Ophyra aenescens predation study.
o

no
Comparisons of cephaloskeletons of Ophyra aenescens and Musca
domestica. It was determined by Keilin and Tate (1930) that a correla¬
tion exists between certain anatomical structures of fly larvae and
their modes of life. In order to gain further knowledge of the feeding
habits of 0. aenescens and M. domestica larvae through the use of ana¬
tomical evidence, the cephaloskeletons of third-instar larvae were
dissected and mounted on microscope slides in balsam.
Areas of the cephaloskeletons compared (Figure 21) were the ventral
surface of the basal sclerite(B.) and the oral sclerite(0.).
Results. The basal sclerites of M. domestica and 0. aenescens are
shown in Figures 22 and 23 respectively. Note the presence of longi¬
tudinal ridges(L.R.) in the basal sclerites of both species. In Figure
22, the salivary duct(S.D.) can be seen where it passes into the
intermediate seleri te(I.).
The oral sclerites of M. domestica are shown in Figure 24. They
consist of two lateral hooks(L.H.), nearly uniform in length, joined
ventrally by a median ventral arc(M.V.A.). Both the median ventral arc
and the lateral hooks are shrouded by the remains of the cuticle. Oral
grooves(0.G.) leading to the oral aperature can be clearly seen.
The oral sclerites of 0. aenescens (Figure 25) consist of two
lateral hooks of unequal lengths. The left hook, shown protruding above
the right hook, is the shorter of the two. It fits into a groove on
the right hook which enables both hooks to work together. Both hooks
are joined ventrally by a median ventral arc.
In addition to the oral sclerites, accessory oral sclerites are
present in the bucco-pharyngea1 armature of 0. aenescens larvae (Figure
25). Beneath the lateral hooks are two oral bars(0.B.). The anterio-

111
Figure 21 .
Areas on the cepha1oskeieton of Ophyra
aenescsns compared with those of Musca
dones tica.

í»»
112
Figure 22. Basal seler i te of Musca domestica showing
longitudinal ridges(L.R.) and salivary
duct (S.D.).
-igure 23.
Da sa i se ¡ er i te of Opiv.-va aeYiescens show i r.g
longitudinal ri aces (L.P.).

I i 3
Figure 2b. Oral sclerites of Musca domestica showing the
intermediate soler ite(I.), the lateral hooks
(L.H.), the median ventral arc (M.V.A.), and
the oral grooves (0.G .) •
Figure 25. Oral sclerites of Cvhyra aer,escens shewing the
lateral hooks(L.H.), the median ventral arc
(M.V.A.), the oral bars(0.B.), the anterior
ribfaon(A.R.), and the cutaneous teeth(C.T-).

114
dorsal borders of the oral bars articulate with two additional sclerites
that join anterior to the tips of the lateral hooks to form the anterior
ribbon(A.R.)• Beneath each oral bar is a row of cutaneous teeth(C.T.),
but only one row is visible in Figure 25-
The longitudinal ridges in the basal sclerites of both species are
indicative of saprophagous behavior and the accessory oral sclerites and
unequal lateral hooks of 0. aenesoens are indicative of predaceous
behavior (Keilin and Tate, 1930). The combination of characteristics,
i.e. the longitudinal ridges and the oral sclerites, indicates that
larvae of 0. aenesoens can live either in a saprophagous manner or they
can be carnivorous.
Competition Studies with Hermetia illucens
Competition studies were performed in the laboratory to gain know¬
ledge of interactions that could occur in the field if the habitats of
H. illucens and those of three other flies, M. domestioa, 0. aenesoens,
and S. rob'iista, overlapped. Experimental designs and larval diets are
shown in Tables 15 and 16. Larvae were grown in 360-ml plastic cups
containing 200 ml of diet. Each cup represented a replication, and
each treatment was replicated four times. Treatments consisting of only
one species of fly were used as controls. After larvae or eggs were
added, cups were covered with screen and placed in the growth chamber.
Trials were terminated when adults of M. domestica, 0. aenesoens,
and S. robusta emerged in their respective control groups. Viable larvae
of H. illucens were counted when experiments ended. Since the soldier
fly larvae tended to stay at the bottom of the cups, they were easily
located. The remaining medium was floated to separate adults from
pupae of the competing fly species when applicable.

115
Table 15- Experimental designs of and the larval diets used in the
competition studies involving larvae of Eermetia illucens
vs. larvae of Musca domestica and Ophyra aenescens.
Treatment No.
T reatment
1
10a
0.
aenescens
, 1
1arvae
D
2
25
0.
aenescens
larvae
3
10
R.
illucens
1 arvae
4
25
H.
illucens
1 a rvae
5
10
0.
aenescens
larvae
+ 10 R. illucens larvae
6
25
0.
aenescens
larvae
+ 10 3. illv.cens larvae
7
10
G.
aenescens
1arvae
+ 25 H. illucens larvae
8
10
M.
domestica
eggs
9
25
M.
domestica
eggs
10
50
M.
domestica
eggs
11
10
H.
illucens
1arvae
12
25
H.
i 1lucens
larvae
13
50
H.
illucens
1arvae
14
10
M.
domestica
eggs +
10 H. illucens
1arvae
15
25
M.
domestica
eggs +
1 0 H. illucens
1 arvae
16
25
M.
domestica
eggs +
25 H. illucens
larvae
17
25
M.
domestica
eggs +
50 H. illucens
larvae
18
50
M.
domestica
eggs +
10 H. illucens
1 a rvae
TM
Larval Diet: 5 parts CSMA + 3 parts water
Represents the number of larvae or eggs per replication with four
replications per treatment.
° Larvae of 0. aenescens were first instars; larvae of H. illucens
were from 1 to 1.5 cm in length.

116
Table 16. Experimental design of and the larval diet used in the compe¬
tition study involving larvae of Rermetia illucens and larvae
of Sarcophaga robusta.
Treatment No. Treatment
1
10a
H.
illucens
i b
larvae
2
10
S.
robusta
larvae
3
25
S.
robusta
larvae
4
10
S.
robusta
1 arvae + 1 0 H.
illucens
larvae
5
25
S.
robusta
1 arvae +10 H.
illucens
1 arvae
Larva 1
Diet: 200 ml CSMAâ„¢
400 ml fish mea 1
kkO ml water
ca. 800 ml very moist medium
Represents the number of larvae per replication with four
replications per treatment.
k Larvae of S. robusta were first instars; larvae of H. 'illucens
were from 1 to 1.5 cm in length.

117
Results of studies with Hevmetia illucens and Q-phyva aenescens•
When the number of 0. aenescens larvae per replication was equal to or
less than the number of h. illucens larvae, the larval mortality of
0. aenescens increased significantly, but the mortality of E. illucens
larvae was not significantly affected (Table 17). When larvae of 0.
aenescens outnumbered those of H. illucens, the larval mortality of
0. aenescens was not affected, but the mortality of H. illucens was
increased by a nonsignificant amount. This indicates that the larval
development of 0. aenescens can be suppressed, but not prevented, by
the presence of H. illucens in the larval medium. This has not been
demonstrated in the field.
Results of studies with Eemetia illucens and Eusca domestica-
Larvae of H. illucens were not able to completely prevent the develop¬
ment of larvae of M. domestica when the two species were reared together
(Table 18). The mortality of M. domestica was increased to 90.0 and
86.0% when 25 larvae were reared with 25 and 50 larvae of H. illucens
respectively. These were significant increases over control mortality.
Control mortality was high because house fly eggs were used instead
of first-instar larvae, but at the time, additional larvae of H. illucens
were not available and the experiment could not be repeated.
The ability of these two species to develop in the same containers
during this experiment, but not during others, may have been due to
variables such as container size, number of larvae per unit area of
medium, and moisture content of the larval medium. Larvae of H. illucens
tended to remain on the bottoms of the cups of medium, perhaps the
wettest location in the cups. House fly larvae tended to remain in the
upper two thirds of the cups of medium, perhaps seeking drier areas than

Table 17. Results of the competition study between Herrnetia illuoens and Ophyra aenescens.
Tr ealinent s
Reps
1
2
3
ll
5
6
7
10 0.
a
aenesaens
25 0. b
aeneucens
10 //.
illuoens
25 I!.
illuoens
10 0.
aeneaaenc
10 //.
ill noons
25 o. +
aiinéscens
10 H.
illuoeno
10 0.
+
aufitdoaena
25 II.
illuoens
1
10
25
10
25
7
9
24
10
i
25
2
7
23
10
2 4
/
10
19
9
6
25
3
9
20
10
21
6
10
24
10
7
25
'i
8
18
10
23
8
10
19
9
a
25
X
8.5XYC
21.5
10.0
23.3
7.0 V
9.8
21.5
9.5
7.0Y
25.0
X Mortality
15.0
14.0
0.0
7.0
30.0
2.5
14.0
5.0
3o.o
0.0
Indicates number of 0. aeneevews larvae used in each rep.
Indicates number of II. illuoens larvae used in each rep.
Represents mean number of viable individuals at the end of the experiment.
Note: Means having common letters are significantly different (p < .05).

Table 18. Results of the competition study between Hermstia illucens and Musca domestica
Rrps
n
9
10
11 enliuents
1 1
12
13
16
10 M.
domen L ica
25 M.
dome plica
50 M.
ci. /mao l iaa
10 //. ,
•11
illucenc
25
i lluooim
50 II.
i l luacna
10 II. (
domp.u L ica
10 II.
illuoetvi
1
8C
7
21
I 0
25
i>9
2
8
2
7
13
9
10
2't
58
8
10
3
5
3
1(3
10
23
50
2
10
5
6
15
18
10
21
'll
7
9
X
6.3C
9 - 6YZ
16.5
10.0
23. J
57.0
5.8
93
% Moit a 111 y
37-5
62.0YZ
67.0
0.0
7.0
6.0
52.5
7.5

Table 18.
Cont¡nued.
1 re ;UnieM Is
Reps
15
ir>
17
10
25 M.
dome atiaa
f 10 II.
illuaena
25 M.
domestica.
+ 25 li.
illuaena
2 5 M.
domed iaa
+ 50 //.
illuaena
50 n.
domestica
+ 10 II.
i l luoeiiB
1
12
10
3
2'i
5
5/
19
10
2
0
to
1
23
5
50
1 3
10
3
6
10
3
25
2
50
21
10
6
10
3
25
2
58
11
10
X
0.0
10.0
2.5Y
25.3
3.5Z
57.0
16.0
10.0
% Morta1 Ity
68.0
0.0
90. 0
3-0
06.0
5.5
60.0
0.0
a I ml i entes t lie number of M. dome ótica eggs used In each rep.
Indicates the number of //. illucp.no larvae used in each rep.
° Represents the mean number of viable Individuals at the end of the experiment.
Note: Pairs of means having common letters are significantly different (p < 05).

121
3. illucens. Thus, the two species developed in separate locations in
the same cups with little chance of coming in contact.
A situation such as this was seen in the field at the tilling site
(Figure 31). Chips had been added to very wet manure which, when tilled,
was ca. 29 cm above the level of the walks. A definite moisture gradient
existed in the chips-manure mixture. Deep in the manure, at the level
of the walks, there was a large population of soldier flies. Near the
top of the manure-chips mixture, 2 to 4 cm beneath the surface, was a
large house fly population. The populations of the two species were
separated vertically by 25 to 27 cm and neither could be detected with¬
out digging into the manure. This was an unusual situation created by
the depth of the manure-chips mixture and the moisture gradient within.
Results of studies with Hermetic illucens and Sarcophaga robusta.
In treatments 4 and 5, where both species were reared together, the
mortality of S. robusta larvae was increased significantly, but the
mortality of 3. illucens was not significantly affected (Table 19).
Since the larvae of S. robusta require only 3 days to complete
their larval development, the reduction in their population by 3.
illucens was not expected. Because larvae of S. robusta and 3. illucens
prefer media similar in moisture content, it is possible that the
bottoms of the cups of medium were the sites preferred by both fly
species. Sarcophaga robusta may have been unable to compete success¬
fully with 3. illucens for the preferred sites and was displaced in
numbers reflected by the increases in mortality shown in Table 19.

122
Table 19. Results of competition study between Hermetia iZZucens and
Sarcophaga robusta.
Treatments
Reps
1
2
3
4
5
10 H.
iZZucens
10 s. b
robusta
25 S. 10 S.
robusta robusta
10 H.
iZZucens
25 5.
robusta
10 H.
iZZucens
1
10C
1
24
4
10
12
10
2
10
8
21
2
9
12
10
3
10
8
23
2
10
12
10
4
10
8
19
2
1 0
8
10
z
40
31
87
10
39
44
40
X
C,
10.0
7.8X
21 . 8Y
2.5X
9.8
1 1.0Y
10.0
/o
Mortality 0.0
22.5
13.0
75.0
2.5
56.0
0.0
a
1 ndj cates
number of
H. iZZucens larvae used in
each repl¡cation.
b
1 ndicates
number of
S. robusta
larvae
used in
each rep 1¡cation.
c
Represents
the number of viabl
e individuaIs at
the end
of the
experiment.
Mote: Pairs
of means
(x) having
common
1etters are signi
f icantly
different (p < .05).

123
Insect Growth Regulators (iGR's) and Organophosphorus Larvicides
Feed-through studies with dimilin and methoprene. Methoprene
(Zoeconâ„¢ ZR-515 10% tech.) and dimilin (Thompson-Hayward TH 6040 25%
tech.) were added to poultry feed to produce diets of the following
concentrations:
IGR
Amount added to
136.4 kg of basal
layer diet(g)
Concentration
of treatment
diet(ppm)
TH 6040
5.44
10
TH 6040
0.54
1
ZR-515
13.60
10
ZR-515
1.36
1
Diets were fed to treatment groups of 44 week-old Babcock B-300 laying
hens randomized in range houses measuring 2.4 by 4.6 m (Figure 26). The
10-ppm diets were fed for 107 days after which the 1-ppm diets were fed
for 14 days. One week elapsed between the change of IGR treatment
1 eve 1s.
Manure was bioassayed three times while 10-ppm diets were fed, and
twice while 1-ppm diets were being fed. Samples were seeded with eggs
of M. domestica. Pupal traps were set twice while the 10-ppm diets
were being fed.
Resu1ts. Feed consumption of hens fed methoprene and dimilin is
shown in Table 20. At both dietary levels, birds eating the dimilin
diets consumed feed in amounts that were significantly greater than
those consumed by birds in the control groups. Birds eating methoprene
diets consumed less feed that the birds in their respective control
groups, but the differences were not significant.
During sampling period 9 (Table 20), hens were inadvertently
allowed to run out of feed, which accounts for the consumption level of
only 77.18 g/bird per day.

124
1—1
3
2
»—!
3
2
2
1
3
2
1
“7
D
Treatment Group Diet
1 Control
2 ZR-515
3 TH 6040
Figure 26. Assignment of diets containing ZR-515 and TH 6040 to
treatment groups in range houses.

125
Table 20. Average daily feed consumption (g/bird per day) of hens fed
diets containing TH 6040 and ZR-515 at 1 and 10 ppm.
Perioda
ZR-515
Growth Regulator
TH 6040
Control
Dietary Level -- 10
ppm
1
106.69
108.96
104.42
2
111.23
113.50
104.42
3
102.15
118.04
102.15
4
104.42
113.50
97.61
5
113.50
122.58
111.23
6
118.04
127.12
118.04
7
113.50
131.66
111.23
8
104.42
122.58
108.96
9
77.18
127.12
106.69
X
105.68A
120.56B
107.19A
Dietary Level -- 1 ppm
1
106.69
111.23
106.69
2
106.69
124.85
111.23
X
106.69C
118.04D
108.96c
Average daily feed consumption was calculated at the end of
each 12-day period.
Mote: Mean values in the same row without common letters are
significantly different (p < 0.01) according to Duncan's Multiple
Range Test.

126
There were no significant differences in egg production due to diet
at the 10-ppm level (Table 21), but the birds consuming the dimilin diet
were laying at a numerically higher level than the other two groups.
At the 1-ppm level, hens consuming the methoprene diet were laying
significantly fewer eggs than the hens in the control groups. This was
attributed to the hens being allowed to run out of feed as mentioned
above. There were no significant differences in the production levels
of the dimilin and control groups (Table 21), but the birds in the
dimilin group maintained a higher numerical production level.
Results from the manure bioassay are shown in Table 22. At both
dietary 1GR levels, fly mortality was significantly greater than mor¬
tality in the control groups. At the 10-ppm level, methoprene and dimilin
produced fly mortalities of 85.00 and 74.17% respectively. At 1 ppm,
methoprene and dimilin produced fly mortalities of only 17.5% which is
too low to be of any practical value.
The data from the pupal traps were nonconclus ive (Table 23) since
mortality in all three groups was nonsignificant. In the control group,
46.48% of the pupae failed to eclose for some unexplainable reason.
The wood chips may have become contaminated with a toxic substance
prior to use in the pupal traps. When Abbott's formula was used to
correct the mortalities, only 16.11 and 12.80% of the mortalities was
due to methoprene and dimilin respectively. These low mortalities
may have been due to the rapid breakdown of the IGR's plus the lack of
larval contact with the IGR's in the manure pack.
Topical application of methoprene. Experiment 1 — The experi¬
mental design used at the tilling site is shown in Figure 27. Blocks 2

127
Table 21. Average hen-day product i on (?á) of hens fed diets containing
TH 6040 and ZR-515 at 1 and 10 ppm.
Growth Regulator
Period3
ZR—515
TH 6040
Con tro 1
Dietary
•
Level -- 10 ppm
1
51.35
50.39
52.41
2
41.49
36.95
31.73
3
53.10
52.76
46.07
4
61.66
73.30
66.14
5
37-36
65.88
65.17
X
49.09A
55.96A
52.30A
Dietary
Level -- 1 ppm
1
26.94B
62.560
57-87C
a Average
hen-day production was
calculated at the
end of each
21-day period.
Note: Mean values in the same row without common letters are
significantly different (p < 0.01) according to Duncan's Multiple
Range Test.

128
Table 22.
Bloassay of
ZR-515 at 1
manure from
and 10 ppm.
hens
fed diets
containing TH 6040 and
Growth Regulator
Samp 1e
No.
ZR-515
TH 6040
Contro 1
Di
i etary
Level --
1 0 ppm
1
- 30a
, b
1/10
6/10
10/10
2
- 30
1/10
7/10
8/10
3
- 30
2/10
7/10
10/10
4
- 30
1/10
6/10
10/10
5
- 92
1/10
1/10
10/10
6
- 92
1/10
2/10
10/10
7
- 92
0/10
1/10
9/10
8
- 92
1/10
1/10
10/10
9
- 107
3/10
0/10
9/10
10
- 107
3/10
0/10
10/10
11
- 107
3/10
0/1 0
10/10
12
- 107
2/10
0/10
10/10
1
18/120
31/120
116/120
X
1.50A
2.58A
9.67B
% Morta1ity
85.00
74.14
3.33
Dietary
Level --
1 ppm
1
- 11
1 0/10
8/10
10/10
2
- 11
10/10
6/10
9/10
3
- 11
10/10
9/10
10/10
4
- 11
9/10
7/10
10/10
5
- 14
6/10
9/10
10/10

129
Table 22. Continued.
Sample No.
ZR-515
Growth Regulator
TH 6040
Control
Dietary Level -- 1
ppm
6 - 14
7/10
8/10
9/10
7 - 14
8/10
10/10
8/10
8 - 14
6/10
9/10
8/10
z
66/80
66/8o
74/80
X
8.25C
8.25C
9.25C
% Morta1ity
17-50
17.50
7.50
The numbers to the right of the hyphen indicate the day of the
experiment on which the sample was collected, e.g. sample 1 was
collected on day 30 of the experiment.
k Represents the number of adult house flies that formed when
10 eggs were set.
Note: Mean values in the same row without common letters are
significantly different (p < 0.05).

130
Table 23. Data from pupal traps set in manure from hens fed diets
containing TH 6040 and ZR-515 at 10 ppm.
Replication
ZR-515
Growth Regulator
TH 6040
Control
1
1/6a
6/16
7/13
2
10/20
11/27
16/28
3
2/8
3/11
12/21
4
9/15
15/21
3/9
Y.
22/49
35/75
38/71
% Mortality
55.10
53-33
46.48
3 Represents
the number of
pupae that eclosed <
over the number of
pupae found in the trap.

131
1.3m
Check
B1 ank
—
538 mg/m2
B1 ank
1078 mg/m2
Block No,
1
2
3
4
5
gure 27. Experimental design for testing the effects of ZR-515
sand granules on larval populations of Musca domestica
in the manure collection area of a poultry house.

132
and 4 were left untreated to prevent overlap of treatments. Methoprene
(Zoecon 0.4% tech.) sand granules were applied at rates of 538 and
1076 mg/m2. The weights of granules applied to blocks 3 and 5 (Figure
27) were 2.08 and 4.17 g respectively.
Immediately after treatments were applied and 3 days later, samples
were collected for laboratory bioassay. Each sample was seeded with 25
eggs of M. domestica.
Resu1ts. Mortality produced by methoprene sand granules at
1076 mg/m2 was significantly higher than the mortalities in the other
two groups (Table 24). When corrected by Abbott's formula, mortalities
produced by the 1076 and 538 mg/m2 treatments were 94.3 and 42.9%
respectively.
In the second group of samples, there were significant differences
in the mortalities produced by all three treatments (Table 24). When
corrected by Abbott's formula, mortalities produced by the 1076 and
538 mg/m2 treatments were 95.82 and 62.91% respectively.
Results indicate that methoprene sand granules are effective for
at least 3 days post-treatment. When the granules are applied at
1076 mg/m2, fly mortality of more than 90% can be expected. Field
tests, however, may not produce similar results as indicated by the
feed-through trials.
Few pupae were formed in the sample replicates of manure treated
with 1076 mg/m2 of methoprene, because at this high concentration, the
1arvae were killed.
Experiment 2 -- At the University of Florida Poultry Science Farm,
methoprene (Zoecon 0.4% tech.) sand granules were sprinkled on manure
beneath two treatment blocks of 10 hens caged in pairs. Weights of the

133
Table 24. Number of pupae, number of pupae eclosed, and per cent
mortality when larvae of Musca domestica were reared in
poultry manure containing two levels of methoprene sand
granu les.
Samp 1e
No.
1076 mg/m2
Treatments
538 mg/m2
Check
No.
Pupae
No.
Eclosed
No.
Pupae
No.
Eclosed
No.
Pupae
No.
Eclosed
1
oa
0
7
4
5
3
0
0
18
4
10
3
0
0
16
7
J
6
6
7
2
14
9
25
23
E
7
2
55
20
55
35
X
0.5A
5.0B
8.8B
% Morta 1 i ty*3
98.0
30.0
65.0
2
24
1
22
16
21
O 1
I
0
0
14
3
16
16
0
0
21
4
18
15
0
0
30
4
28
27
E
24
1
87
27
OO
WJ
79
X
0.3c
6.8d
19-8E
% Morta1ity
99.0
78.0
24.0
Represents the number of pupae formed from 25 house fly eggs.
k Equals the number of eclosed pupae subtracted from 100.
Note: Mean values in the same row without common letters are
significantly different (p < 0.01).

134
granules used for the 1076 and 538 mg/m2 treatments were 0.57 and 0.29 g
respectively. Samples were collected as for experiment 1 above.
ResuIts. In the first group of samples, house fly mortalities in
groups treated with methoprene were significantly higher than the check
mortalities (Table 25). When corrected by Abbott's formula, mortalities
produced by the 1076 and 538 mg/m2 treatments were 93-9 and 97.0%
respectively.
In the second group of samples, mortalities in groups treated with
methoprene were significantly higher than the check mortalities, despite
the high check mortalities. When corrected by Abbott's formula, mor¬
talities produced by the 1076 and 538 mg/m2 treatments were 70.6 and
88.2% respectively.
Results indicate that methoprene sand granules are effective for at
least 3 days post-treatment. When the granules are applied at 1076 mg/m2,
mortalities greater than 90% can be expected, at least on the first day.
Results from field tests may differ from laboratory tests as previously
stated. It cannot be explained why the lower concentration of methoprene
produced numerically higher mortalities in both sample groups. Lack of
pupae formation in the 538 mg/m2 treatment in the second group of samples
indicated the presence of a high level of methoprene, possibly due to
collection of manure from an area where nonuniform application occurred.
High check mortalities in both experiments were attributed to the
use of house fly eggs in the manure samples. House fly eggs hatch
better in wet media and media in this experiment were somewhat dry.
TM
Laboratory studies with CGA 72662. CGA 72662 (Ciba-Geigy 10%
soluble concentrate) was added to the diets of six species of flies.
The fly species and diets utilized are shown in Table 26.

135
Table 25. Number of pupae, number of pupae eclosed, and per cent
mortality when larvae of Musca domestica were reared in
poultry manure containing two levels of methoprene sand
granules that were applied at the University of Florida
Poultry Science Farm.
Samp 1e
No.
1076 mg/m2
T reatments
538 mg/m2
Check
No.
Pupae
No.
Eclosed
No.
Pupae
No.
Eclosed
No.
Pupae
No.
Ec1osed
1
1 4 3
1
18
0
21
21
9
1
24
1
12
12
11
0
22
1
14
14
12
2
20
0
19
19
E
46
4
84
2
66
66
X
1 .0A
0.5A
16.5B
% Morta1ityk
96.0
98.0
34.0
2
5
5
1
1
14
13
12
0
0
0
4
4
9
3
2
2
1 0
10
8
2
1
1
8
7
y
2-
34
10
4
4
36
34
X
2.5C
1 .OC
8.50
% Morta1ity
90.0
96.0
66.0
3 Represents the number of pupae formed from 25
house fly eggs.
k Equals
the number of eclosed pupae subtracted
from 100
.
Note: Mean values in the same row without common letters are
significantly different (p < 0.05).

136
Table 26. Fly species and diets used for CGA 72662 laboratory studies.
Fly
D iet
M.
domestica
350
ml
CSMAâ„¢
10
ml
fish mea f
200
m 1
test solution or H20
H.
iliucens
500
m 1
CSMAâ„¢
300
ml
test solution or H20
S.
robusta
100
ml
CSMAâ„¢
200
ml
fish mea 1
220
ml
test solution or H20
P.
regina and
150
ml
CSMAâ„¢
F.
canicuiaris
100
m 1
f i sh mea 1
160
m 1
test solution or H20
0.
a&nescens
400
ml
CSMAâ„¢
100
ml
horn fly dry mix3
275
ml
test solution or H20
3 960g wheat flour,
720g fish mea 1
l,
120g Na2C03, 400g alfalfa meal
(Greer, 1975).

137
Resul ts. Results are summarized in Table 27 and the raw data are
in Appendix 2. At the concentrations tested, CGA 72662 produced 100%
larval mortality in all flies except H. iltucens and F. canicularist
and 100% pupal mortality in H. illucens. Flies other than H. iVluoens
and F. cani.aula.in-s produced no pupae. No larval remains could be found
in the growth media which indicates early larval death.
Concentrations of CGA 72662 were within the range of concentrations
to be used in the field. Under field conditions, concentrations may
become diluted after being applied to fresh poultry manure. Therefore,
the concentrations of CGA 72662 used in this experiment were much higher
than field populations of flies would be likely to encounter.
Larval house fly dosage-mortality curve for CGA 72662. The test
concentrations of CGA 72662 added to house fly larval diets in this
experiment were 1.0, 0.75, 0.50, 0.25, and 0.10 ppm. The diet used is
shown in Table 26.
Results. The resulting mortality data are shown in Table 23. Two
trials viere performed and data were combined. In one trial, the number
of larviform pupae found at different concentrations of CGA 72662 was
recorded (Table 28). The lowest concentration producing larviform
pupae was 0.25 ppm. All larviform pupae formed at this concentration
eclosed. Those produced at higher concentrations did not eclose. At
the highest concentration of CGA 72662, 1.0 ppm, only five uneclosed
larviform pupae formed in the four treatment replications. It was
therefore assumed that other larvae at this and higher concentrations
died prior to reaching the pupal stage.
A larviform pupa formed during the experiment is shown in Figure 28.
Larviform pupae were comparable in size to third-instar larvae and
slightly darker in color than norma 11y-formed pupae.

Table 27. Summary of per cent larval mortality in CGA 72662 laboratory studies.
Fly Species
Control
0. 10
CGA 72662 Concentrations^)
0.075 0.06 0.05 0.0'i
0.03
0.025
M. domestica
27.5
—
—
100.0
100.0
100.0
100.0
—
H. illucens3
o
o
100.0
87-5
—
85.0
—
—
52.5
S. robusta
7.5
100.0
100.0
—
100.0
—
—
100.0
0. aenescens
20.0
100.0
1 00.0
—
100.0
—
—
100.0
F. canicularisa
25.0
77.5
57.5
—
75.0
—
—
52.5
P. regina
5-0
100.0
100.0
—
100.0
—
—
100.0
Values for H. illucens and F. canicularis represent percentages of pupae that formed. Pupal
mortality was 100% in all cases except controls. Other fly species died in the larval stages.

Table 28. Larval mortality and larviform pupae formation resulting from various levels of CGA 72662 in
growth media of house flies.
T reatment
(ppm)
2
3
4
5
6
1.0
0.75
0.50
0.25
0.10
Control
15/15a
14/15 (H)b
13/15
do)b
3/15
<2)b
0/15
2/15
15/15 (5)b
15/15 (4)
13/15
(13)
4/15
(3)
5/15
3/15
15/15
15/15 (13)
13/15
(7)
2/15
(D
2/15
1/15
15/15
15/15 (9)
11/15
(a)
3/19
(3)
1/15
2/15
14/15
8/15
7/15
4/15
3/15
0/15
12/15
8/15
6/15
3/16
1/15
0/15
13/15
11/15
5/15
2/16
2/15
0/15
13/15
9/15
7/15
4/15
1/15
1/16
z
112/120
95/120
75/120
25/126
15/120
9/121
X Mortality
93.33
79-17
62.50
19.84
12.50
7-44
Corrected
% Morta1ityC
92.80
77.49
59.49
13-40
5.47
a Represents:
initial no.
of 1st instar
larvae -
no. of
ec1osed
pupae
or adults
initial no
. of 1st
instar
1 arvae
Represents number of iarviform pupae found.
„ , ■ a,i . . r i % mortality in treatment - % mortality check v .
Corrected using Abbott's formula: 10b' - V ;7^T~')Tr~7~hT:A-T X 100
% morta1ity check
V.D

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

The probability, log dose, upper and lower fiducial limits, and
probits are shown in Table 29. The probit curve with the LCS0 of
0.45 ppm is shown in Figure 29. The slope of the probit curve indicates
relative heterogeneity in the susceptibility of the laboratory house
fly population to CGA 72662 (Hoskins and Gordon, 1956).
Comparison of the efficacies of CGA 72662 and commercially used
organophosphorus larvicides. CGA 72662 and three commercially used
organophosphorus larvicides were applied to manure in one house at the
tilling site. Sectioning of the manure collection area, assignment of
treatments, and the application rates of the compounds are shown in
Table 30. Treatment blocks measured 7-6 m along the walk by 1.2 m wide.
The procedures for mixing the test concentrations are given in Table 31.
Time interval for the collection of samples was as follows:
Sample
No.
1
2
3
4
5
6
7
Time
1 nterva1
pretreatment, but on treatment day
4 days post-treatment
7 days post-treatment
11 days post-treatment
14 days post-treatment
28 days post-treatment
35 days post-treatment
Sampling ceased after the 'larval population mean of a treatment block
exceeded that of the control block. All sampling ceased 35 days post¬
treatment because manure in the control block became too dry to support
fly populations.
Results. The larval population means for all treatments during
each sampling period are shown in Table 32 and Figure 30. The active
period for the larvicides is shown in Table 33.

i 42
Table 29-
The probabilities, log doses, upper and
and probits from the probit analysis of
mortality data.
lower fiducial limits,
CGA 72662 dosage-
3 s
p
ñ-:;a
•) ■:
5
-
L QW
c. R
¡ j -j p
c
p
Pr
331
1
3
r* t
_
1 1
•
37
- 1 L
, 77
- 1 1
9
1 G
2
. 6 7
0
r*. -p
-
1 1
•
ri
- 1 1
* 57
- 1 0
'J c
c.
• Py
3
•j
- 3G
-
11
„
11
- 1 l
.45
1 \j
•33
~7
1 T>
i.*.
0
• 0 4
-
11
•
03
- 1 1
. 35
- 1 0
0 1
P
• 25
,3
.75
-
1 0
♦
9 7
-1 1
.23
- 1 0
7 6
. 38
p
n
• Go
-
10
»
9.2
- 1 l
. 21
- t 0
7 2
3
. 4 3
G
r. ~7
-
1
•
n 7
- 1 i
.16
- 1 0
ó 3
-7
- •">
• >
Cl
C
. 7 4
—
1 â– 
5 3
- 1 1
. 1 0
-1C
í 4
-_p
.53
• 0 ?
-
15
7 9
- 1 l
. 0 6
- 1 0
6 1
"7
♦ 66
i J
. 1 G
-
i
•
7 0
- 1 l
.0 2
- 1 0
-j 3
p
â– 7 T?
• i
[ l
0
• 1 5
-
1 0
•
0 L
- 10
.64
-1 0
4 6
3
. 6
n 0
L
0
» 20
-
1 0
•
59
- 1 3
. 7 0
- 1 0
36
¿+
.16
1 :
Q
• 2 T>
—
1 0
•
49
- 1 0
.53
- 1 0
27
4
♦ —»0
L 4
3
• 0
• J J
-
L 0
31
- 1 0
. 47
- 1 0
20
4
, 4 H
i 3
c
• 3 5
-
1 0
•
2 3
- 10
.33
- 1 0
i 3
't
.61
1
Q
* 0
-
1 0
n
115
-10
.20
-1 0
0 ó
4
.75
1 7
0
♦ 4-3
-
1 3
•
3 3
- 1 9
. 2 0
_ t;
9 6
4
. 8 7
L 0
0
• 30
-
1 V
•
0 1
- 1 j
. 1 1
-9
^ 2
5
. 0 0
1 9
. 35
- 9
#
93
1
.03
-9
L> Zj
. 1 3
?. 7
0
• 60
- 0
•
36
— O
.95
— 9
73
p
♦ 4. “3
«c i
,3
♦ 65
_o
•
73
— 9
-37
—y
7 0
p
. 39
V.'
-7 -V
♦ / • i
— 0
•
73
— 9
.73
— 9
20 *
> k
77
0 CT )
—
♦ 7 5
— <■)
*
9 1
-9
. 7 0
— 9
5 1
0
♦ c 7
>
3
. 3 )
♦
-j 1
— Q
. 0 0
- y
40
0
n b 4
•1. Ty
.-» 6 Tj
- O
•
-9
. 50
— 0
27
.6
< >' 4
j r.
3
Q P;
—
•
â– ? yt
-0
. 37
- 0
1 \j
-.0
• 23
3
* * }
— 9
•
22
— 9
.34
-9
j 6
ó
• 34
_ . j
0
0 92
•
1 3
-9
. 3 1
- 4
'j 1
J
• 4 1
7 ‘7
pi
• 93
-0
9
1 4
—9
. 27
— 0
9 ó
♦ 4 B
J 0
3
> '' 4
— 0
9
l 3
— 0
.23
- s
9 0
* 5 5
p ?
♦ j
- 9
â– *
94
-9
. 19
- j
3 4
• 6*^
—• o
* 0 6
. j
â– *
3 3
—0
. 1 3
— 0
7 6
• T'
j 3
1 j
# Q”7
- 3
♦
~ 1
-9
. 0 7
— 0
V
• 1
9 -
3 -
j
- 0 3
— 6
•
5 'J
—6
.98
— 0
3 5
* . 'To
3 ;.S
. 3 9
. j.
•
64
g
• 35
— 8
3 5
7
» ...

Probits
Figure 29. Probit curve, fiducial limits, LCso, and regression equation for CGA 72662 dosage-
morta1 ity data.

1 44
Table 30. Sectioning of the manure collection area, assignment of
treatments, and the application rates of CGA 72662 and
the organophosphorus larvicides applied to poultry manure.
T reatment
blocks in
manure Treatment Application Rate
col 1ection
area
dimethoate
d i chlorvos
dich1orvos
D TM
Ravap
CGA 72662 (0.05%
CGA 72662 (0.05%
1.0 gal/100 ft2
0.5 gai/100 ft2
1.0 ga1/100 ft"
1.0 gal/100 ft2
1.0 gal/100 ft2
0.5 gal/100 ft2
(3.84 1/9.12 m2)
(1.92 1/9-12 m2)
(3.84 1/9.12 m2)
(3.84 1/9.12 m2)
(3.84 1/9.12 m2)
(1.92 1/9.12 m2)
Cont ro1
CGA 72662 (0.]%)
CGA 72662 (0.1%)
1.0 gal/100 ft2
0.5 gal/100 ft2
(3-84 1/9-12 m2)
(1.92 1/9-12 m2)

145
Table 31- Mixing the test concentrations of CGA 72662 and the
organophosphorus larvicides applied to poultry manure.
Contents of Test Concentrations
Compound
Name
Formu1 ation
[% EC)
Compound
(ml)
H20
(mi.)
Test
Concent ration{%)
dimethoate
23.4
192.0
3840
1 .0
d i ch1orvos
23.4
76.8
3840
0.5
D TM
Ravap
23 + 5.7
153.6
3840
1.0
CGA 72662
1 0
20.0
3840
0.05
CGA 72662
10
40.0
3840
0.1

Table 32. Larval population means for all treatments during each
sampling period when poultry manure was treated with
CGA 72662 and three organophosphorus larvicides.
T reatments
dimethoate
dich1orvos
D tm
Ra vap
Samp 1ing
Period
Time
I nterva1
(days) 1
1.92 1/9.12m2 3.
2
84 1/9.12m2
3
4
1
0
355.25A
235.00C
386.00A
305.00B
2
4
53-OOA
13.00D
19.00C
20.00C
3
7
335.25B
421.00A
3 5 - 2 5 B
269-OOC
4
11
465.50A
302.50B
527.25A
33.50C
5
14
6
28
7
35
Note: For each sampling period, treatment means without common
letters are significantly different (p < .05) using the method of
J. W. Tukey.

147
Table 32. Extended.
T reatments
CGA 72662 (0.05%) Control CGA .72662 (0.1%)
3.84 1/9.12m2 1.92 1/9.12m2 3-84 l/9-12m2 1.92 l/9-12m2
5
6
7
8
9
266.00C
261.50C
104.75D
117.25D
119.000
10.000
27.00B
20.50C
0.50E
0.25E
0. OOF
0. OOF
108.250
0. OOF
0. OOF
0.00C
3.75C
62.25C
0.00c
O.OOC
0.00B
62.25A
73-75A
0.00B
O.OOB
6.25B
54.75A
55-75A
O.OOB
O.OOB
0.00B
103.50A
107.75A
O.OOB
O.OOB

LARVAi. ¡ iEANS
Sampling Period (day of treatment)
Figure 30. Larval population means for all treatments during each sampling period when poultry
manure was treated with CGA 72662 and three organophosphorus larvicides.

Table 33. Larvicidal activity period of compounds tested in the CGA 72662 organophosphorus larvicide study.
f rea Unen t
d imethoate dichlorvos Ravapâ„¢ __ CGA 72662 (0.05%) Control CGA 72662 (0, It)
1.92 1/9.12m2 3.86 1/9.12m2 3.86 1/9.12m2 1.92 1/9.12m2 3.86 1/9.12m2 1.92 1/9.12m2
I 2 3 6 5 6 7 8 9
1 0
2 6
T i me
Samp liny Interval
Period (Days)
3 7
6 I 1
5 16
6 28
7 35

150
At the beginning of the experiment, there were significant differ¬
ences between larval population means in the treatment blocks, but the
control block had the lowest mean of all blocks (Figure 30).
By the second sampling period, only blocks treated with dimethoate
and CGA 72662 (0.05%) at 1.92 1/9-12 m2 had larval population means
that were significantly greater than that of the control block (Table
32, Figure 30). There were no significant differences between the
larval population means of blocks treated with dichlorvos at 3.85 1/
9.12 m2 and Ravap, and the larval population mean of the control block.
Larval means of all other blocks were significantly lower than the
control block.
By the end of the third sampling period, fly resurgence had begun
in the blocks treated with dimethoate, dichlorvos at 1.92 1/9-12 m2,
and Ravap, and the larval population means were all significantly
greater than the larval population mean of the control group (Figure 30).
Larval means of all other treatment blocks were significantly lower than
the control block population. Larval populations in the blocks treated
with both rates of CGA 72662 were reduced to zero.
At the end of the fourth sampling period, populations in blocks
treated with dimethoate and dichlorvos at 1.92 1/9.12 m2 continued to
resurge, and resurgence also began in the block treated with dichlorvos
at 3-84 1/9.12 m2 (Figure 30). Sampling was discontinued in these
blocks. The block treated with Ravap shewed a decrease in larval popu¬
lation, but by the end of the fifth sampling period, the population was
considered too numerous to count. Therefore, the larval population
counted during the fourth sampling period was the last recorded count
and sampling in the Ravap-treated block also ceased.

151
For the remainder of the experiment, there were no significant
differences between the larval populations in the block treated with
the lower level of CGA 72662 (0.05%) at 1.92 1/9.12 m2 and the control
block (Table 32, Figure 30). Populations in other blocks still being
monitored were reduced to zero.
The activity period for dimethoate, dichlorvos at 1.92 1/9.12 m2,
and Ravap was 7 days (Table 33). The activity periods for dichlorvos
at 3-84 1/9.12 m2 and CGA 72662 (0.05%) at 1.92 1/9.12 m2 were 11 and
14 days respectively. CGA 72662 (0.05%) at 3-84 1/9-12 m" and both
rates of 0.1% CGA 72662 were still active 35 days after treatment was
applied.
Larvicidal activity of CGA 72662 when tilled into wet manure.
Manure from poultry houses at the tilling site was cleaned out leaving
the bottoms of the manure collection areas ca. 14 cm below the surface
of the walks. At this depth, the tines of the tiller were too short
to reach the fresh manure being dropped. This, in conjunction with
heavy rains and flooding, resulted in the rapid development of large
populations of S. illucens.
To give consistency to the manure and to raise it to a level that
the tiller tines could reach, wood shavings were added to the manure
collection areas as previously described. The manure was tilled once,
leaving the surface of the manure-wood shavings mixture ca. 29 cm
above the surface of the walks.
Afterward, manure was not tilled for 1 week. During this time,
large populations of house flies developed near the surface of the
manure-wood shavings mixture, along with the soldier fly populations
further beneath the surface (Figure 31).

11.5" (29.21 cm)
5.5" (13.97 cm)
Bottom of Manure
Collection area
Water Level
Manure-Wood Shavings
Mixture
A - HOUSE FLY POPULATIONS
o - SOLDIER FLY POPULATIONS
gure 31. Cross-section of manure-wood shavings mixture 1 week after tilling, showing relative
locations of house fly and soldier fly populations.
VJ~)
ho

153
A section of the manure collection area of one house was divided
into four blocks and treatments were assigned as shown in Figure 32.
A 0.1% solution of CGA 72662 was mixed as shown beiow:
40 ml of CGA 72662 + 3.84 1 H2O = 0.1% solution
The application rate was 3-84 1/9.12 m2. Treatment blocks 2 and 3 were
tilled twice weekly and blocks 0 and 1 were left untilled.
Pretreatment samples were collected on the day the experiment began
and additional samples were collected at weekly intervals. Larvae of
house flies, soldier flies, and little house flies were counted as in
the previous experiment. Sampling was discontinued after 5 weeks because
of climatic conditions.
Results. Larval means from each treatment block for each sampling
period are shown in Table 34.
The pretreatment house fly samples revealed significant differences
between treatment-block population means. Blocks to be treated with
CGA 72662 had numerically lower populations than the remaining two blocks.
One week after treatment, there were no significant differences
between the house fly population means in the four blocks, but in the
untilled block sprayed with CGA 72662, the house fly population mean
was 2.75 as compared to 37-50 in the tilled CGA 72662 block (Table 34).
Manure was so wet that house fly larvae were pupating just below the
surface of the manure instead of at the edges of the manure pack. in
the untilled GCA 72662 block, fly larvae coming up to pupate were being
exposed to the concentrated CGA 72662 on the manure surface. In the
tilled CGA 72662 block, CGA 72662 may have been diluted by the tilling
and was slower acting.

Tilled
Twice
Weekly
Mot
Tilled
154
gure 32. Treatment area, assignment of treatments, and tilling
schedule in the CGA 72662 tilling trial.

155
Table 34. Weekly
house
tilled
treatment means
fly populations
twice weekly.
of house fl
from manure
y, soldier f1
treated with
y, and 1 i ttle
CGA 72662 and
_ a
T reatments
Week
0
1
2
3
Musca domestica
0 = Precount
97-OOB
187.50A
136.00AB
143.00AB
1
2.75A
39.50A
17.50A
37-50A
2
0.00c
47.50A
22.753
0.00c
3
0.00B
16.25A
16.25A
0.00B
4
93-50A
31.25B
35.253
0.00c
5
209.00B
376.25A
150.25C
0.00c
Hevmetia
illucens
0 = Precount
31-25A
1 • 75 B
2.25B
0.50B
1
18.50B
7.50BC
12.00AB
1 .ooc
2
0.00B
0.75B
7-75A
0.00B
3
0.00B
8.50A
6.00AB
0.00B
4
0.00B
15.25A
9.25A
0.00B
5
0.00B
39-OOA
12.75A3
0.00B
Fannia canicularis
0 = Precount
10.00A
28.50A
27.50A
11.00A
1
0.00A
2.25A
1 .50A
2.00A
0
0.00B
3.25AB
7.00A
0.00B
3
0.00A
0.25A
0.00A
O.OOA
4
0.50A
0.00A
0.00A
0.00A
5
0.00A
0.00A
0.00A
O.OOA
3 Treatment 0 = CGA 72662, no tilling; Treatment 1 = control, no
tilling; Treatment 2 = control, tilled twice weekly; Treatment 3 =
CGA 72662, tilled twice weekly.
Mote: Means in the same row having the same letter are not
significantly different (p < .05).

156
By the second week after treatment, house fly populations in both
blocks treated with CGA 72662 were reduced to zero. Population means
in the tilled control were significantly lower than the population means
in the untilled control.
By the fourth week post-treatment, house flies began to resurge in
the untilled block treated with CGA 72662 and the larval population mean
was significantly higher than the population means in either control
block. Larval populations in the tilled CGA 72662 block remained at
zero.
At the end of the fifth week, house fly resurgence had occurred
in all treatment blocks except the tilled CGA 72662 block (Table 3^,
Figure 33) .
Initial H. illucens populations were much lower than the house fly
populations (Table 3*0- By the second week, soldier fly populations
were reduced to zero in blocks treated with CGA 72662 and populations
in both untreated controls were greater than populations in the CGA
72662 blocks. This situation continued through the end of the experi¬
ment (Table 3^, Figure 3^).
There were no significant differences in the little house fly
population means at the beginning of the experiment, but by the second
week, populations in both CGA 72662 blocks had been reduced to zero.
By the third week there were no significant differences between little
house fly population means. This situation remained the same through
the end of the experiment (Table 3^+, Figure 35).
CGA 72662 added to drinking water as an oral iarvicide. The test
concentrations of CGA 72662 utilized were 5-0, 1.5, 10.0, and 20.0 ppm.
The treatment and sample collection schedule is shown in Table 35.

House Fly Larval Means
Figure 33. Weekly treatment means of house fly populations from manure treated with CGA 72662
and tilled twice weekly.
U~1

Week
Figure 3^. Weekly treatment means of soldier fly populations from manure treated with CGA 72662
and tilled twice weekly.
CO

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

160
Table 35•
Treatment and sample collection schedule when CGA 72662 was
added to the drinking water of laying hens as an oral
1 arv icide.
Day 1
Began daily treatment with CGA 72662 at
1 . 5 and 5•0 ppm.
Day 7
First sample group of 5 cups of manure
per treatment was collected.
Day 11
Second sample group of 10 cups of manure
per treatment was collected.
Day 1 6
Discontinued daily treatment with CGA
72662 at 1.5 and 5-0 ppm. All birds were
returned to 500 ml H2O 3 times daily.
Day 22
Began daily treatment with CGA 72662 at
10 and 20 ppm. Replaced polyethylene
under cages and in water troughs.
Day 29
First sample group of 5 cups of manure
per treatment was collected.
Day 30
Discontinued daily treatment with CGA
72662 at 10 and 20 ppm. All birds were
returned to 500 ml H20 3 times daily.
Replaced polyethylene under cages.
Day 33
One sample group of 5 cups of manure per
treatment was collected 3 days post¬
treatment.
Day 35
One sample group of 5 cups of manure per
treatment was collected 5 days post¬
treatment .

161
ResuIts. Both levels of CGA 72662 significantly increased house
fly mortality in the first trial (Table 36). The mortalities of the
5.0-and 1.5-ppm treatments were 87.56 and 80.89% respectively, and
corrected mortalities were 84.85 and 76.72% respectively.
The results of the second trial are shown in Table 37- Manure from
both CGA 72662 treatments produced house fly mortalities significantly
greater than the control mortality. Treatment levels of 20.0 and 10.0 ppm
produced mortalities of 100.00 and 92.00% respectively, and corrected
mortalities of 100.00 and 91.11% respectively.
Mortalities produced by manure from treated hens 3 days post¬
treatment were still significantly greater than mortality rates in
control groups (Table 38). Manure from hens formerly receiving CGA
72662 at levels of 20.0 and 10.0 ppm produced mortalities of 99-33 and
80.00% respectively. Corrected mortalities were 98.97 and 69.39%
respect ively.
Results of samples bioassayed 5 days post-treatment are shown in
Table 39- There were no significant differences in mortality due to
treatment and, after correction, mortalities produced by manure from
both former treatment groups were less than 10%.
Blacklight Electrocutor Grid Traps for Adult Fly Surveys
Fly catches from experimental blacklight electrocutor grid traps
(Danalco"^) were processed for 6 months. Species of flies observed
includes M. domestica, H. illucens, S. calcitrans, E. irvitans, and
Ophyra s p.
Results. The monthly catches of the above species of flies in the
two traps are shown in Table 40. In nearly all cases, trap A caught
greater numbers of flies than trap B. Except for April and September,

162
Table 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.
20 ppm
1 0 ppm
Control
15/15
•
14/15
2/15
15/15
13/15
1/15
15/15
14/15
3/20
15/15
14/15
1/15
15/15
14/15
1/15
Tota 1 Morta1ity
75/75
69/75
8/80
Mean Mortality
15.00
13-80
1 .60b
% Mortality
100.00
92.00
10.00
Corrected % Mortality3
100.00
91.11
—
Corrected using Abbott's formula:
% treatment mortality - % check mortality ^ ^ qq
100 - % check mortality
Significantly lower than other two treatments at p < 0.05%.

163
Table 37- Mortality of immature house flies
hens collected when CGA 72662 was
water at the rates of 1.5 and 5.0
in the manure of
added to the dri
ppm.
laying
nk i ng
5.0 ppm
1.5 ppm
Control
13/15
6/15
4/15
12/15
8/15
3/15
10/15
11/15
3/16
11/15
7/15
4/16
11/15
8/15
0/17
14/15
14/15
0/15
13/15
13/15
7/15
13/15
14/15
2/15
14/15
15/15
1/15
15/15
15/15
3/15
14/15
15/15
2/15
13/15
13/15
4/15
15/15
15/15
2/15
14/15
15/15
6/16
15/15
13/15
0/15
Tota 1 Morta1ity
197/225
182/225
41/229
Mean Morta1ity
13.13A
12.13A
2.73B
% Morta1ity
87-56
80.89
17.90
Corrected % Mortality3
34.85
76.72
—
3 Corrected using Abbott's formula:
; treatment mortality - % check mortality 1Q0
100 - % check mortality
Note: Means having different letters are significantly
different (p < 0.05)*

164
Table 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.
20 ppm
1 0 ppm
Control
15/15
13/15
6/15
15/15
10/15
5/15
15/15
12/15
2/15
15/15
11/15
5/15
15/15
13/15
5/15
15/15
12/15
5/15
14/15
13/15
12/15
15/15
13/15
6/15
15/15
11/15
4/15
15/15
12/15
2/1 5
Tota 1 Morta1ity
149/150
120/150
52/150
Mean Mortality
14.90A
12.00B
5.20C
% Mortality
93.33
80.00
34.67
Corrected % Mortality3
98.97
69-39
—
Corrected using Abbott's formula:
; treatment mortality - % check mortality y
100 - % check mortality
Note: Means having different letters are significantly
different (p < 0.05).

165
Table 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.
20 ppm
1 0 ppm
Control
4/15
3/15
0/15
1/15
2/15
2/15
2/15
0/15
1/15
2/15
1/15
0/15
0/15
1/15
0/15
Tota 1 Morta1ity
9/75
7/75
3/75
Mean Morta1ity
1.80
1.40
0.60
% Morta1ity
12.00
9.33
4.00
Corrected % Mortality3
8.33
5.55
—
3 Corrected using
% treatment mortality -
Abbott's formula:
â–  % check morta1ity v
100
100 - % check
morta1ity

1 66
Table 40. Monthly catches of Musca domestica, Hevmetia illucens,
Stomoxys calcitvans, Kematobia irvitans, and Ophyra sp.
in two blackllght electrocutor grid traps.
T rap
Apr
May
Months
Jun Jul
. Aug
Sep
M. dome
stica
A
21110
34380
67880
124930
68070
18460
B
11 570
21470
14200
48940
26360
6930
H. illucens
A
4o
140
180
1 40
90
10
B
50
100
20
100
50
50
S. calcitvans
A
950
1250
720
40
10
260
B
640
890
210
60
0
120
H. irvitans
A
60
30
0
0
0
0
B
40
20
0
10
0
0
Ophyra
sp.
A
200
770
250
180
20
0
B
220
530
60
60
10
0

167
trap A caught more soldier flies than trap B. Trap B caught greater
numbers of S. oalai-tvans in July than trap A, but trap A caught the
most S. aaloitrcms during the other months. There were slight numerical
differences in catches of H. ivvitans , but trap A caught more than trap
B except during July. Except for the month of April, trap A caught more
Ophyra sp. than trap B.
Cow pastures bordered the tilling site on two sides. This accounts
for the numbers of S. calaitrans and H. irritans caught in the traps.
Catches of 3. HZucens in April, May, and June probably consisted
of adults that emerged from pupae formed the previous fall. Subsequent
catches most likely consisted of adults that emerged during April and
May.
Trap catches indicate relatively high populations of Ophyra during
April, May, and June, but adults were never noted at the tilling site
except in the traps.
House fly populations were low in April, peaked in July, and
dropped again in September (Figure 36). The two traps presented two
different views of house fly activity during the 6-month period. This
indicates the necessity of using traps that are uniform when using
trap data for survey work. The low June catch by trap B cannot be
expiained.
Efficacies of Granular Fly Baits
Laboratory and field testing were performed on granular baits pro¬
vided by two different manufacturers. Both groups of tests will be
described separately.

Figure 36. Fluctuation iri house fly populations as recorded by two blacklight traps at the
tilling site.
oo

169
Farnam baits. I he following baits were submitted by Farnam:
Kill'em Fly Killer II (Bomyl) - sand base
Kill'em Fly Bait (Bomyl) - corn cob base
Kill'em Fly Killer (Vapona + ronnel)
SX-70 Fly Bait (methomyl)
Baits were subjected to knockdown and residual tests. The amount of
each bait used was 5 g.
Knockdown test. Results are shown in Table 41 . Kill'em Fly Killer
had the fastest knockdown, killing all flies in 10 min. Kill'em Fly Bait
killed all fiies within 1 hr and 40 min, and SX-70 Fly Bait and Kill'em
Fly Killer M killed all flies in 4 hr and 40 min, and 4 hr and 50 min
respectively.
Residual test. Results are shown in Table 42. By the 10th day of
testing, all baits had picked up moisture from the air. SX-70 Fly Bait
had the longest residual, still producing a mortality rate of greater
than 25% after a 6-week testing period. Testing of Kill'em Fly Killer
ceased on day 10 after the mortality rate dropped to 20% and testing of
Kill'em Fly Killer ¡I and Kill'em Fly Bait ceased on day 19 after mor¬
tality rates dropped to 30 and 10% respectively.
Although testing was discontinued on one bait on day 10 and two
baits on day 19, these baits were maintained through the end of the
experiment. As a check, flies were exposed to the discontinued baits
once every week. Mortality never reached the last recorded values in
Table 42 and continued to drop with time.
The sudden fluctuation in mortality rates produced by SX-70 Fly
Bait during days 15 through 33 is unexplainable. Equally unexplainable
is the 22.5% control mortality on day 12.

Table 41. Results of knockdown test with Farnarn baits.
8nl u
I I me
InlervsI
(I) KlII'fM My Killer ••,H
(BuriyIj Jund line)
(7) KHl'cm My Polt,M
(B «¡¡y I) (corn cob hotel
Renlirntlon _ . 'rnllfollnn ,
Pei cent ' Ter cent
A B C 0 Hof In Illy A * C ft flor loll ty
ID Kill'd r It Klll-r'"
(VitpuuH ♦ rotmel)
Ri'iil I mi Ion
ARCO
Ter cent
Mor t ñ111 y
ID $« m n, n.n'M
("i* I licoiy I f _
fti'pl Ini Inn
A B C 3
Per cent
Mor Iw11ly
0*
0
0
0
0*'
o or
0
0
0
0
0 0 0000 00
0
0
0
0
0.0
I
1
0
0
0
7 S
5
2
7
6
37.5 lo 10 10 10 100.0
0
2
J
I
15.0
2
1
7
4
I
75 0
9
6
10
9
f*5 o
3
4
6
l
35.0
3
I
5
4
7
15 o
9
R
10
10
92.5
J
4
6
2
37 5
4
1
5
4
J
I/O
9
B
10
10
97 5
6
5
A
8
62.5
5
J
5
7
3
45 0
10
9
10
10
97 5
6
5
6
fl
67 5
4
4
5
7
4
50.0
10
9
10
10
97 5
6
5
6
R
67 5
1
5
7
7
5
6o.o
10
9
10
10
975
7
6
6
B
6/5
8
5
7
7
5
6o.o
10
9
10
10
97 5
R
6
6
R
70.0
9
5
7
7
5
60 0
10
9
10
10
97 5
6
6
7
8
77.5
10
5
7
7
6
67 5
10
10
10
10
100.0
B
6
7
0
77.5
II
5
7
e
6
65 0
8
6
7
R
77.5
12
5
7
8
6
65.0
8
6
7
R
77 5
II
5
7
8
6
65.0
R
6
7
9
75.0
14
5
8
8
6
67.5
0
6
7
9
75 0
15
5
8
R
6
6/5
8
6
7
9
75 0
16
5
8
R
6
675
8
7
7
9
77 5
12
5
9
9
6
72.5
R
8
9
9
85 0
18
5
9
9
6
77.5
8
8
9
9
85.0
19
5
9
9
6
77 5
R
8
9
9
R50
70
5
9
10
7
775
ft
R
9
9
RS 0
71
6
9
10
7
R0 0
n
R
9
9
05 0
22
6
9
10
8
87.5
0
R
9
9
05.0
2)
7
9
10
8
05.0
R
8
9
9
05.0
24
10
9
10
9
95.0
9
9
10
9
97.5
75
10
9
• 0
9
950
9
9
10
9
92.5
76
10
9
10
10
97.5
9
9
10
9
97 5
71
10
9
10
10
975
9
9
10
10
95 0
78
10
9
10
10
97.5
9
9
10
10
95 0
79
10
10
10
10
100.0
10
10
in
10
100 0
Cnch number refrenen!* « IO-m|n Interval.
^ fitch number rrpirtmlt the number of filet Hill’d n*il of » totnl of 10.
c _ . , number dr*«J „ ,n_
Per cent mortality • - I* 100
Hotel All filet user! In this e*perln*c"t wri e five • labnretury rolrmy
Hie* were tened while «net the!I red with €0».

Table ^2. Results of residual tests with Farnam baits.
iih?
(I) Kill*'* Ply Mller™ (?) Klll'cm fly Balt™ U) Kill'pm My Killer™ ('#) SX 70 My P*»t ™ Control Control
lüfü’íií i,a"d ^ Urxnjf I ^ ^coin rob bese£ (Vapono * roitMpJ) (me t Itomy I f with sugar watei
I Ime
Application
fer cent
R>’pl lent Ion
Per cent
Aef
1 1
a* Inn
I'er cent
Re|
11
el Inn
Per cent
Aep
1
at Inn
Per
ent
Aep 1 I
a t 1 on
Per cent
Inter val
A
e
c
0
Morml11 y
A
A
c
D
Mnr1 allt y
A
0
C
0
Mollullty
A
A
C
0
Urn t a 1 1 t y
A
A
c
0
Mm t a 111 y
A
A
c
D
Mm tal 11 y
l"
10
lo
10
./
ioo.oc
10
10
10
10
It'll 0
10
lo
10
lo
lon.o
10
in
10
10
100 0
0
0
0
0
0
0
0
0
0
0
0.0
2
10
10
10
10
100.0
10
10
lo
10
t HO 0
10
lo
lo
10
100 0
10
10
10
in
loo 0
0
0
0
0
0
0
n
0
0
0
0.0
1
10
9
10
9
95.0
10
10
10
10
lon.o
10
10
10
II)
100.0
10
9
10
10
975
0
0
0
0
0
0
0
0
0
0
0.0
b
10
10
10
1
92 5
lo
9
10
1
90 0
lo
9
10
10
97 5
10
10
10
10
ino.o
0
0
0
0
0
0
0
0
0
0
0.0
5
9
10
9
9
92 5
9
9
9
9
90.0
0
5
0
7
70.0
10
10
10
in
loo.o
0
0
0
0
0
0
0
0
0
0
0.0
10
9
10
10
10
9/ 5
9
9
9
9
90.0
J
0
i
b
70.0
10
10
10
10
100.0
0
0
0
0
0
0
0
0
0
0
0.0
11
10
10
6
9
97.5
lo
10
10
10
100 0
10
lo
10
10
100,0
0
0
0
0
0
0
1
0
7
0
75
17
10
1
10
9
90.0
8
10
lo
10
95.0
10
10
10
10
100.0
1
0
0
0
7
5
3
3
3
0
77 5
1*
6
6
70.0
8
7
75 0
A
9
-
B5.0
0
0
0
0
0
0
-
0.0
• A
9
10
10
9
95 0
8
6
5
9
750
9
9
9
9
90 0
0
0
n
0
0
0
0
0
1
7
7.5
16
8
8
9
9
65.0
8
9
8
9
05 0
9
9
10
9
92 5
0
0
0
0
0
0
0
0
0
0
0.0
19
1
1
1
I
)0.0
1
0
1
0
loo
10
9
A
9
9b 7
0
0
0
0
0
0
0
0
0
0
0.0
27
10
1C
10
10
loo o
0
0
0
0
0
0
0
0
0
0
0.0
n
10
10
10
10
ion o
0
0
0
n
0
0
o
0
0
0
0.0
7b
A
6
1
10
01 A
0
0
0
0
0
0
0
0
0
0
0 9
15
10
7
7
9
91 7
0
0
0
1
2
5
0
0
0
2
50
7t
7
9
5
i
71.7
3
b
b
b
17
5
0
2
0
0
5 o
79
7
6
A
6
9M
0
0
0
0
0
0
0
0
0
0
0.0
II
1
6
b
1
bf.J
0
0
0
0
0
0
0
n
0
0
0.0
J7
6
7
1
10
92 J
0
0
0
0
0
0
0
0
0
0
0.0
u
7
9
A
9
9b 3
0
0
0
0
n
0
0
I
0
0
2.5
16
7
9
9
A
91-7
0
0
0
0
0
0
0
0
0
0
0 0
17
9
5
A
9
86 1
0
0
0
0
0
ft
0
0
0
0
0.0
)6
7
6
7
A
73 1
0
0
0
0
0
0
0
0
0
0
0.0
19
1
b
7
3
20 9
0
0
0
0
0
0
0
0
0
0
0.0
bo
b
b
5
b
bft 0
0
0
0
0
0
0
0
0
0
0
0.0
b|
5
3
7
2
II. o
0
0
0
0
0
0
0
0
0
0
0 0
mnbeM liHlcst* weekend* or IwrlldayS-
a total of 10.
femala tonta file», Niiflra «lire’af (t . ), from a laboratory colony.
F»rh nrmber ifpr»tent? a one-day 11 irvo Interval. Skipped
*’ Each number representa the mirnhct of 11 lea killed not of
c number dead „
Ter cent mmtolliy X 100
Hoto» All filet used In Ihls experiment rceia five-day old
files were sexed while anesthetized with COj.

172
Field test. The baits described above plus Golden Malrin without
Muscamone were tested in the field with and without the addition of
Lure'em !l attractant supplied by Farnam. Golden Malrin with Muscamone
was used as a standard. When baits were tested alone, 5 g were used.
When tested with the attractant, 5 g of bait and 3 g of the attractant
were used together. The Lure'em II attractant was never used in the
same bait station with Golden Malrin with Muscamone.
Results. Treatment means by treatment are shown in Table 43 and
Figure 37- Treatment means by sex are shown in Table 44 and Figure 38.
Raw data is in Appendix 3. Kill'em Fly Killer II plus Lure'em II
attractant killed a significantly greater number of flies than the
other treatments including the Golden Malrin with Muscamone standard
(Table 43, Figure 37). Treatment means for Kill'em Fly Killer II plus
Lure'em II attractant and Kill'em Fly Bait plus Lure'em i! attractant
were 158.25 and 116.13 flies per bait station respectively. All other
means were less than 30.0 flies per bait station (Table 43).
Lure'em II attractant improved the kills of all baits except SX-70
Fly Bait. Golden Malrin without Muscamone plus Lure'em II attractant
killed a significantly greater number of flies than the Golden Malrin
with Muscamone standard. Kills produced by the standard bait were
numerically lower than all other kills except those of Kill'em Fly
Killer (Table 43).
Kill'em Fly Killer killed males and females of M. domestica in
equal numbers (Table 44, Figure 38), but all other baits killed more
females than males. Three baits killed significantly greater numbers
of females than males. Lure'em II attractant increased the catches of
females for all baits except SX-70 Fly Bait. Golden Malrin without

173
Table 43. Treatment means by treatment in Farnam bait field trial.
T reatment
No.
Treatment
Treatment
Mean
1
+
5
Kill*em Fly Killer IIâ„¢ + Lure'em
IIâ„¢
158.25
A
2
+
5
Kill'em Fly Ba i tâ„¢ + Lure'em 11
116.13
B
4
SX-70 Fly Baitâ„¢
28.63
C
4
+
r*
:>
SX-70 Fly Bait + Lure'em II
20.13
CD
6
+
5
Golden Malrinâ„¢ w/o Muscamoneâ„¢ +
Lure'em II
20.00
CDE
3
+
5
TM
Kill'em Fly Killer + Lure'em 11
19.63
DEF
6
Golden Malrin w/o Muscamone
15.75
DEFG
2
Kill'em Fly Bait
11.38
FGH
1
Kill'em Fly Killer ¡1
8.00
GHI
7
Golden Malrin with Muscamone
7.88
GHI
7
Golden Malrin with Muscamone
6.13
HI
3
Kill'em Fly Killer
0.50
1
Note: Treatment means without common letters are significantly
different (p < 0.05).

160
CO
<
LU
>-
_l
u_
LU
CO
128
96
64
32
0
o
o
1
o
JjL
1+5
2
2+5
3
3+5
4
4+5
6
6+5
7
o
.TM
K¡ 1 1 'em Fly Ki Her I I™
Kill'em Fly Killer il + Lure'em II
Ki I 1 ' em Fly Ba i tâ„¢
Kill'em Fly Bait + Lure'em II
Ki 1 l'em Fly Ki 1 lerâ„¢
Kill'em Fly Killer + Lure'em II
SX-70 Fly Baitâ„¢
SX-70 Fly Bait + Lure'em II ^
Golden Mai rinâ„¢ w/o Muscamone
Golden Mairin w/o Muscamone + Lure'em II
Golden Malrin with Muscamone
O
o
o
1+5 2 2+5 3 3+5 9 4+5
Treatment
6+5 7
Figure 37. Farnam bait field trial treatment means.
4T-

175
Table 44. Treatment means by sex in Farnam bait field trial.
T reatment
No.
T reatment
Sex
T reatment
Mean
1
Kill1em Fly Killer 1!^^
F
9.25'
M
6.25"
1
+ 5
Kill'em Fly Killer II + Lure'em IIâ„¢
F
231.ooT
M
85.507
2
Ki111em Fly Baitâ„¢
F
12.25
M
10.50
2
+ 5
Kill'em Fly Bait + Lure'em II
F
157.50
M
74.75
3
Ki 1 1 ' em Fly Ki 1 1 erâ„¢
F
0.50
M
0.50
3
+ 5
Kill'em Fly Killer + Lure'em II
F
31.75
M
7-50
4
SX-70 Fly Baitâ„¢
F
31.75
M
25.50
4
+ 5
SX-70 Fly Bait + Lure'em II
F
*ic
' o
LA
O
CA
M
9.75"
6
TM TM
Golden Malrin w/o Muscamone
F
18.50
M
13.00
6
+ 5
Golden Malrin w/o Muscamone + Lure'em II
F
30.75
M
9.25
7
Golden Ma1rin with Muscamone
F
9.25
M
3.00
7
Golden Malrin with Muscamone
F
9.25
M
6.50
Indicates significant difference between sexes (p < 0.05).
Indicates significant difference between sexes (p < 0.01).

250
200
150
100
50
0
T~
~r â– 
1
T~
1 1
j-
r
i ii r 1 1
O
1
Kill 'em Fly Killer II
A
1+5
Kill'em Fly Killer II + Lure'em II
O
U
2
Ki 1 1 1 em Fly Ba i t^
_
2+5
Kill'em Fly Bait + Lure'em 11
3
Ki 1 1 'em Fly Ki 1 lerâ„¢
3+5
Kill'em Fly Killer + Lure'em 11
h
SX-70
Fly BaitTM
4+5
6
SX-70
Golden
Fly Bait + Lure'em II
Malrin^11 w/o Muscamone
6+5
Golden
Mairin w/o Muscamone + Lure'em II
O
7
Golden
Mairin with Muscamone
o -
$
o —
o’
-
O
-
O
-
o
O
o
O
V
o
<>
&
1
1—
o-
0
C
1
O
i
a , i_ Í
66+57 7
1 1+5 2 2+5 3 3+5 4 4+5
Treatment
Farnam bait field trial treatment means by sex,
o>

177
Muscamone plus Lure'em II attractant caught 3 times as many females as
Golden Malrin with Muscamone.
Burroughs Wellcome bait. The bait submitted by Burroughs Wellcome
•was Atroban (BW 21Z) 0.25% sugar bait. Golden Malrin with Muscamone
was used as a standard. These two baits were subjected to knockdown,
residual, attractancy, and field tests. The amount of each bait used
per replication was 5 g.
Knockdown test. Both baits required the same amount of time to
produce mortality rates of 100% (Table 45). Atroban took much longer
to actually kill flies than Golden Malrin with Muscamone, but after
60 min, flies exposed to Atroban were completely incapacitated. However,
criterion for death was total lack of movement, and flies could not be
considered dead until all movement ceased.
Residual test. Results are shown in Table 46. After 6 weeks of
testing, Atroban and Golden Malrin with Muscamone baits were producing
mortality rates of 87.5 and 100.0% respectively. Mortality was still
occurring in the manner described in the knockdown test.
Attractiveness test. There were no dead flies in the Atroban bait
station (Table 47). This indicates that Atroban was either unattractive
to house flies or, due to the slow killing action of Atroban, flies died
after leaving the bait station.
There were 25 dead flies in the Golden Malrin with Muscamone bait
station.
Field test. Results are shown in Table 48. No flies were found
in the Atroban bait stations, but a mean of 12.5 flies were found in
the Golden Malrin with Muscamone bait stations. The ability of Atroban

Table 45. Results of the knockdown test using BW 21Z and Golden Ha 1rin
TH TM
with Muscamone fly baits.
BW 21 Z
Baits
Gol den Ma1rin
with Muscamone
Cont ro1
T i me
Replication
Per cent
Rep 1¡cation
Per cent
Repl¡cation
Per cent
1n terva1
A
B
C
D
Mortality
A
B
C
D
Mor ta1ity
A
B
c
D
Mortality
oa
0
0
0
oc
o
o
CL
0
0
0
0
0.0
0
0
0
0
0.0
1
0
0
0
0
0.0
2
0
1
4
17-5
0
0
0
0
0.0
2
0
1
0
0
2.5
2
0
1
4
17-5
0
0
0
0
0.0
3
0
1
0
1
5.0
5
0
1
4
25.0
0
0
0
0
0.0
4
0
2
0
1
7.5
5
0
2
4
27.5
0
0
0
0
0.0
5
0
2
0
1
7.5
5
0
3
4
30.0
0
0
0
0
0.0
6
0
2
0
1
7.5e
5
0
3
7
37-5
0
0
0
0
0.0
7
1
2
0
1
10.0
5
0
3
7
37-5
0
0
0
0
0.0
8b
1
4
0
2
17.5
5
0
3
7
37.5
0
0
0
0
0.0
9
4
4
2
3
32.5
6
4
7
7
60.0
0
0
0
0
0.0
10
6
6
5
6
57.5
8
7
7
9
77-5
0
0
0
0
0.0
11
8
8
6
6
70.0
9
7
8
9
82.5
0
0
0
0
0.0
12
9
9
8
8
85.0
9
9
9
10
92.5
0
0
0
0
0.0
''j
03

Table 45. Continued.
Baits
BW 21 Z
Gol den Ma1rin
with Muscamonc
Con tro 1
Time
Rep 1¡cation
Per cent
Rep 1¡cation
Per cent
Rep 1¡cation
Per cent
erva 1
A
B
C
D
Morta1ity
A
B
C
D
Mortality
A
B
c
D
Morta1i
13
10
10
8
8
90.0
9
9
9
10
92.5
0
0
0
0
0.0
14
10
10
9
8
92.5
9
9
10
10
95.0
0
0
0
0
0.0
15
10
10
9
8
92.5
9
10
10
10
97.5
0
0
0
0
0.0
16
10
10
10
10
100.0
10
10
10
10
100.0
0
0
0
0
0.0
Each number represents a 10-min interval.
Each number represents a 30-min interval.
Each number represents the number of fl
number dead
es killed out of 10.
Per cent morta1 i ty =
10
X 100
All flies that were exposed to the BW 21Z bait were at this point either dead or completely
incapacitated; since the criterion for death was not met, per cent mortality was figured at 7-5?.
Note: All flies used in this test were five-day-old female house flies, Musca domestica (L.),
from a laboratory colony. Flies were sexed while anesthetized with CO2.

Table ^6. Results of the residual test using BW 21Z and Golden Ma 1 r i n
T M TM
with Huscamone
fly baits.
Baits
BW 21 Z
Go 1 den Ma1rin
with Muscamone
Control
T i me
Rep 1¡cation
Per cent
Rep 1ica tion
Per cent
Replication
Per cent
1nterva1
A
B
C
D
Mortality
A
B
C
D
Mortal it y
A
B
C
D
Mortali
1a
10
9
10
10b
97.5C
10
10
9
10
97.5
0
0
0
0
0.0
2
10
10
10
10
100.0
10
10
10
10
100.0
3
0
0
0
7.5
3
3
9
10
10
80.0
10
10
10
10
100.0
0
0
0
0
0.0
h
10
7
8
9
85.0
9
9
10
9
92.5
2
0
0
0
O
un
6
5
10
10
10
87.5
10
10
10
10
100.0
0
0
0
0
0.0
Each number represents the number of weeks after the completion of the knockdown test.
b Each number represents the number of flies killed out of 10.
c n . . ,.. number dead v 1A
Per cent morta l i ty = ^ X 1 00
Note: All flies used in this test were five-day-old female house flies, ttusca domestica (L.),
from a laboratory colony. Flies were sexed while anesthetized with CO2.

181
Table 47. Results of the attractiveness test using BW 21Z and Golden
MalrlnTM with Muscamone^M fly baits.
Ba i t
No. Flies
Killed
Per cent
Ki 1 1
BW 21 Z
oa
•
o
o
cr
Golden Ma1rin
with Muscamone
25
12.5
Represents the number of flies killed out of 200.
Per cent kill
number dead
200
X 100.
Note: All flies used in this test were five-day-old female house
flies, Musca domestica (L.), from a laboratory colony. Flies were
sexed while anesthetized with C02-

182
Table 48. Results of the f
with Muscamone^M
i e 1 d
fly
test using
baits.
BW 21Z and Golden Malrin"*"^
Ba i
ts
BW
21 Z
Go 1 den Ha 1rin
wi th Muscamone
Replicate
d*
o
+
cf
?
1
0
0
9
5
2
0
0
“7
/
17
3
0
0
0
2
4
0
0
3
7
5
0
0
—
a
E
0
0
19
31
Ave
0
0
4.75
7.75
a
Bait station was lost.

183
to kill flies in the field was not demonstrated although its efficacy
was adequately proven in the laboratory tests.
Contact Residuals
Adult house fly dosage-mortality curve for permethrin. Concentra¬
tions of permethrin (JFU 5819 25% permethrin EC, ICi Americas, Inc.)
tested and responses are shown in Table 49. The probabilities, probits,
log doses, and upper and lower fiducial limits are shown in Table 50.
The dosage-mortality curve with fiducial limits is shown in Figure 39-
The LD5g for JFU 5819 is 18.0 ppm and the regression equation is
Y = 14.25 + 1.46X.
The high LD50 and the relatively shallow slope indicate that some
amount of resistance to permethrin may already exist in this house fly
strain and that more resistance is developing within the gene pool.
Possible cross-resistance to chlorinated hydrocarbon insecticides may
explain this phenomenon as indicated in the literature (Prasittisuk
and Busvine, 1977).
The resDonses (Table 49) are not well distributed between the
mortality values of 10 and 90%. This is due to the fluctuation in
mortality rates produced by the same test concentrations on different
generations of the house fly colony. After 10 to 15 trials, the set
of data in Table 49 was the most reliable set produced.
Permethrin as a contact residual in the laboratory. Permethrin
(JFU 5021A 2 lb/gal EC, ICI Americas, Inc.) was applied to cement, tin,
and plywood templates to run-off at rates of 0.05 and 0.005%.
Female laboratory colony house flies, 3 to 5 days old, were trans¬
ferred by use of a vacuum system to cylindrical 'window-screen cages
12.5 cm high and 7-0 cm in diameter. Open ends of the cages were placed

184
Table 49. Test concentrations and corresponding responses from the
JFU 5819 laboratory bioassay.
Replicat ion
19
Test Concentrations(ppm)
15 10 5 1
Control
1
6/8a
4/9
3/9
2/10
1/8
2/8
2
4/7
4/8
3/9
3/11
1/8
1/9
3
5/8
5/9
3/8
2/11
2/8
1/9
4
5/8
3/8
4/9
1/11
1/8
2/9
y
¿j
20/31
16/34
13/36
8/43
5/32
6/35
X
5.00
4.00
3-25
2.00
1.25
1.50
7a Mortality
64.52
47.06
37.14
18.60
15.63
17.14
Corrected % Mortality'3
57.18
36.11
24.14
1.72
—
—
3 Represents the number
of fl
¡es killed out
of the total
number
of flies tested.
k Corrected using Abbott's formula:
; treatment mortality - % check mortality x .nn
100 - 7a check mortality

185
Table 50. The probabilities, probits, log doses, and upper and lower
fiducial limits from the probit analysis of JFU 5819 dosage-
mortal ity data.
C3S
PROS
PROBITS
LOGOGSE
LO*Em
UPPER
1
0.0 l
2.67
-7 .94
-11.15
-7. 35
2
0.02
2,95
-7.76
- 10*54
-7. 23
3
0.03
3.12
-7.64
- ro. 16
-7. 16
4
0. 04
3.25
-7.55
-9.37
-7 . 10
5
0.05
3.36
-7.43
-9.63
-7. 06
6
0.96
3 .4o
-7.42
-9.43
-7,10
7
0.07
3.52
-7. 36
-9.2 6
-6.93
3
C .03
3.59
-7.3 1
-9.10
-6. 95
9
0. 09
3.06
-7.27
—3 .9 6
-6.93
1 0
0.10
3.72
-7.23
-3.33
-6.90
1 1
0.15
3.96
- 7.06
-8.29
-6. 73
12
0. 20
4.16
-6 .93
—7 .33
-6.69
13
0.25
4.33
-6.3 1
-7.53
-6. 59
14
0-30
4 .43
-6 . / 1
-7.2 4
-6,49
15
0.35
4.61
-6.61
-6.99
-6.36
16
0.40
4 .75
-6.52
-6.31
-6. 20
1 7
0.45
4.37
-6.43
—ó .67
—ó « CO
1 3
0 » oO
5.00
-6.35
-6.57
-5. 77
1 9
0.55
5.13
— 6.56
-6.43
-5. 52
20
0. 60
5.25
— ó . 1 7
—6.41
-5.25
2 1
0.55
5.39
-6.03
-6.34
-4. 97
ry
t- A.
0 .70
5 .52
-5 .99
-6.27
-4.67
23
0.75
5.67
-5.83
-ó.20
-4.34
¿‘■V
0 «30
5.34
-5.77
-6. 1 3
-3. 98
25
0. 35
5,04
-5.64
—6 «04
-3.54
26
0.90
6.23
-5.47
-5.94
-3.00
27
0.91
6-34
-5.43
-5.9 1
-2. 37
33
0. 92
6.41
-5 .33
-5 .39
-2.7.2
29
0.93
6.43
-5.33
-5,36
-2. 57
3 0
C- 94
Ó .33
-5 .23
—5 «32
-2. 39
3 1
C.95
6. Ó4
-5.22
-5.79
-2,19
"V
w •<—
0 .96
6.75
-5.15
-5. 74
-1.95
33
C- 97
6 .83
— o -06
—5 . 69
— 1,66
3 4
0.9d
7.05
-4.94
-5.62
-1 - 23
35
0.99
7.33
-4.75
-5.50
-0.67

Probits
PPM
Figure 33. Probit curve, fiducial limits, and LC5o for JFU 5819 dosage-mortality data. —
O'N

187
on the templates with four cages on each template and 10 flies per cage.
A cotton ball soaked in a sucrose solution was placed on top of each
cage as a food source. Mortality was recorded after 24 hr. Criterion
for death was total lack of movement. Control templates were processed
like the treated templates except that they were not sprayed with
permethrin.
Testing continued at selected intervals until mortality dropped
below 50%. Between tests, templates were stored outside under the eaves
of the laboratory to simulate aging under field conditions. After each
use, cages were baked in an oven at 148.9 C for a 24-hr period.
Results. Mortality on the tin templates dropped below 50% within
3 days (Table 51) and testing ceased after 8 days. Mortality on the
cement block templates never exceeded 50% and testing was also discon¬
tinued after 8 days (Table 51).
The breakdown of JFU 5021A was slower on wood (Table 51). Mortality
on the 0.005% template began to fluctuate 56 days post-treatment and
dropped below 50% 75 days post-treatment. As a check, the template was
tested again the following summer, but permethrin had broken down
(Table 51) and testing was discontinued.
The 0.05% template continued to produce mortalities of 100% through
1976. When testing resumed in 1977, however, the mortality rate began
to fluctuate (Table 51). Mortality dropped below 50% on 25 August and
8 November, but was back to 77-3% by 5 December. The reasons for this
are not clear, but may be related to lower ambient testing temperature.
The mean mortality for the year was 65.8%.

188
Table 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.
Control 0.05% 0.005%
Date
Morta1ity
Per cent
Morta1ity
Morta1ity
Per cent
Mortal i ty*
Morta1ity
Per cent
Morta1ity
1976
Surface:
Tin
Appl¡cation date:
9/15/76
15 Sep
4/21
19.0
64/102
62.7
60/107
56.1
18 Sep
4/20
20.0
11/47
23-4
22/57
38.6
23 Sep
6/20
30.0
7/60
11.7
6/65
9.2
1976
Su rface:
Cement block
Application date:
9/15/76
15 Sep
3/23
13.0
58/118
49.2
40/88
45.6
18 Sep
4/25
16.0
18/64
28.1
11/47
23.4
23 Sep
1/20
5.0
5/60
8.3
5/53
9.4
1976
Su rface:
Wood
App1ication date:
8/24/76
1 4 Sep
7/65
10.8
45/45
100.0
58/58
100.0
15 Sep
15/122
11.5
113/121
93-4
68/113
60.2
29 Sep
3/50
6.0
53/57
93.0
10/54
18.5
19 Oct
3/31
9.7
37/37
100.0
25/43
58.1
7 Nov
10/80
12.5
81/81
100.0
40/86
46.5
1977
Su rface:
Wood
App1icat ion date:
8/24/76
1 Ju 1
8/49
16.3
41/47
87.2
3/46
6.5
25 Aug
0/46
0.0
23/49
46.9
0/39
0.0
20 Sep
14/62
22.6
36/54
66.7
Testing
Discontinued
11 Oct
5/50
10.0
37/52
71.2
8 Nov
5/51
9.8
28/62
45.2
5 Dec
10/48
20.8
33/45
77.3
X
= 65.8

189
Table 51. Continued.
Date
Control
0.05%
0.005%
Morta1ity
Per cent
Morta1ity
Morta1ity
Per cent
Morta1ity
u _ ,.. Per cent
Mortal 1ty ^ ,..
Mortal 1ty
1978
Surface:
Wood
App1icat ion date:
8/24/76
31 Jan
6/42
14.3
35/42
83.3
28 Feb
0/36
0.0
29/39
74.4
21 Mar
1/39
2.6
26/37
70.8
9 Jun
1/42
2.7
24/48
50.0
6 Ju 1
0/33
0.0
5/40
12.5
3 Aug
1/40
2.5
1/40
2.5

190
Testing continued in 1978. Mortality was 50% on 9 June and dropped
to 2.5% by 3 August. At this point the experiment was terminated.
Permethrin at 0.05% was effective on wood for almost 23 months.
Fluctuation of control mortality throughout the experiment can
only be attributed to an interaction between the flies and the un¬
treated template surfaces.
Synthetic pyrethroids as contact residuals in the field. Three
synthetic pyrethroid compounds, SD 43775 (Shell Development Co., 25% WP
and 10% EC), BW 21Z (Burroughs Wellcome Co., 25% WP and 42.5% EC), and
ICl 143 (1CI Americas, Inc., 5% EC), were applied to wooden panels in
the amounts shown in Table 52.
Several weeks after panels were installed at the tilling site,
there was no evidence to indicate that the compounds were killing flies
as no dead flies were found in the aluminum guttering. In order to
test the efficacy of the compounds, 3“ to 5-day-old female laboratory
colony house flies were processed and exposed to panel surfaces in the
following manner. Flies were sexed while anesthetized with CO2 and
placed 10 to a cage in cages made from 180-ml plastic cups with lids.
Cup bottoms had been removed and replaced with disks of window screen.
In the field, cages were slipped under 23_cm lengths of 3~cm wide
elastic that had been stapled to the panels for the purpose of holding
the cages tightly to the surface. Lids were removed as cages were
secured on the panels. One cage was attached to each panel for a 24-hr
period. Criterion for death was total lack of movement.
ResuIts. All compounds produced mortality rates up to 100% at 121
days post-treatment (Table 53). Knockdown was usually within 15 min
though the onset of death required a much longer period. Flies fluttered

Table 52. Names, formulations, test concentrations, mixing instructions, and application rates of
compounds applied to wooden panels.
Pane 1
No.
Compound
Name
Formu1 ation
and Type
Test
Concentration
(%)
Amount used to
make 500 ml
(g)
Amount applied
to each Side
of Pane 1 (ml)
Appl¡cation
1 ns tructions
C
Control
—
—
—
—
—
1
Shell 43775
25.0% WP
0.05
1 .0
95.0
Appl
y
to
runoff
2
Shell 43775
10.0% EC
0.05
2.5
95.0
Appl
y
to
runoff
3
BW 21 Z
25.0% WP
0.50
10.0
95.0
Appl
y
to
runoff
4
BW 21 Z
25.0% WP
0.25
5.0
95.0
Appl
y
to
runoff
5
BW 21 Z
25.0% WP
0.125
2.5
95.0
Appl
y
to
runoff
6
BW 21 Z
42.5% EC
0.50
5.9
95.0
Appl
y
to
runoff
7
BW 21 Z
42.5% EC
0.25
2.9
95.0
Appl
y
to
runoff
8
BW 21 Z
42.5% EC
0.125
1.5
95.0
Appl
y
to
runoff
9
ICI 143
5.0% EC
0.50
50.0
41.0
Appl
y
@
gal/750 ft2
Note: Compounds were mixed in water.

Table 53. Total and per cent mortality that occurred when 3~ to 5-day-old female house flies were
exposed to synthetic pyrethrolds on wooden panels.
Treatment Number
C
1
2
3
4
5
6
7
8
9
Days Post-treatment:
86
0/10
8/10
6/10
10/10
10/10
10/10
9/10
6/10
7/10
8/10
0/10
9/10
10/10
10/10
1/4
9/10
9/10
9/10
9/10
9/10
1/10
10/10
9/10
10/10
10/10
8/10
10/10
10/10
9/10
9/10
0/10
8/10
6/10
10/10
2/2
10/10
10/10
8/10
10/10
9/10
Total Mortal 1ty
1/40
35/40
31/40
40/40
25/26
37/40
38/40
33/40
35/40
35/40
% Morta11ty
2.5
87.5
77.5
100.0
88.5
92.5
95.0
82.5
87.5
87.5
Days Post-treatment:
100
1/10
10/10
10/10
10/10
10/10
10/10
10/10
9/10
9/10
10/10
0/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
4/10
10/10
10/10
10/10
10/10
8/10
10/10
10/10
10/10
9/10
6/10
9/10
8/10
10/10
9/10
9/10
9/10
1 0/ 1 0
10/10
10/10
Tota 1 Mortal 1ty
1 1/40
39/40
38/40
4o/4o
39/40
37/40
39/40
39/40
39/40
39/40
% Morta11ty
27.5a
97-5
95.0
100.0
97.5
92.5
97.5
97.5
97.5
97.5

Days Post-treatment:
116
8/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
9/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
7/9
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
Tota 1 Mortality
34/39
40/40
40/40
40/40
40/40
40/40
40/40
40/40
40/40
40/40
% Mor t a 1 it y
87.2b
100.0
100.0
100.0
100.0
100.0
100.0
100.0
100.0
100.0
Days Post-treatment:
121
0/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
0/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
0/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
0/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
10/10
Tota 1 Morta1ity
0/40
40/40
40/40
40/40
40/40
40/40
40/40
40/40
40/40
40/40
% Mor ta1ity
0.0
100.0
100.0
100.0
100.0
100.0
100.0
100.0
100.0
100.0
Two replications came in contact with pyrethrin fog when the producer fogged his egg room.
Temperature was ca. 85 F (29.4 C), very hot and very dry.
Note: Treatment numbers are as follows: C = control, 1 & 2 = SD 43775, 3 through 8 = BU 21Z,
9 = 1CI 143.

194
randomly throughout their cages and did not necessarily remain in contact
with the treated surface. No further contact with the panels was needed
to cause the flies to die.
â– x
At 100 days post-treatment, the poultryman fogged one end of his
poultry house while a fly trial was in progress and the mortality rates
of two control panels were affected (Table 53).
When a trial was performed 116 days post-treatment, ambient tempera¬
ture was 29.4 C, but temperatures inside the poultry houses were above
35.0 C. High control mortality during this trial was attributed to the
heat.
When an additional trial was attempted ca. 325 days post-treatment,
a new flock of chickens had been housed on the farm. In between flocks,
houses and panels had been thoroughly sprayed with a disinfectant and
when flies were exposed to the panels, mortality was zero in all cases.
Northern Fowl Mites
Dosage-Mortality Curves for Selected Acaricides
Compounds tested were four synthetic pyrethroids (SBP—1382 (Penick
24.3% EC), BW 21Z (Burroughs Wellcome 42.5% EC), SD 43775 (Shell 10% WDL) ,
and Ectiban (ICI Americas 26.0% EC)], Sev i n (Union Carbide 99-9% crysta-
1ine powder), and malathion (American Cyanamid 96.4% EC).
ResuIts. The test concentrations of acaricides and the total, per
cent, and corrected per cent mite mortality for each concentration tested
are shown in Table 54. Figure 40 shows all six dosage-mortality curves
plotted on one set of axes and the regression equations and LCso's are
shown in Table 55. Probabilities, probits, log doses, and upper and
lower fiducial limits are shown in Appendices 4 through 9-

195
Table 54. Concentrations of acaricides and total, per cent, and
corrected per cent mortality for each concentration
tested against northern fowl mites.
Test Concentration
(ppm)
Tota 1
Mortality
Per cent
Morta11ty
Corrected
% Morta11ty
Penick
1382â„¢
10.00
54/60
90.00
88.10
8.00
54/61
88.52
86.90
6.00
40/62
64.52
58.33
4.00
33/61
62.30
54.76
2.00
25/60
41.67
30.95
1.00
25/62
40.32
28.57
Control
10/61
16.39
—
BW 21 Z
4.25
44/60
73.33
70.99
2.75
33/60
55.00
51 .06
1.75
26/60
43.33
38.36
1.25
19/60
31.67
25.68
1 .00
15/60
25.00
18.43
Control
5/62
8.06
—
Shell SD
43775
10.00
51/60
85.OO
82.74
8.00
45/60
75.00
71.23
6.00
40/60
66.67
61.64
4.00
29/60
48.33
40.53
2.00
21/60
35.00
25.19
1.00
12/61
19.67
7.55
Control
8/61
13.11
—

196
Table 54. Continued.
Test Concentration
(ppm)
Tota 1
Morta1ity
Per cent
Morta1ity
Corrected
% Mortality
TM
1CI Ectiban
6.00
52/60
86.67
85.19
4.00
43/60
71.67
68.52
2.00
34/60
56.67
51.86
1 .00
27/60
45.00
38.89
0.75
21/60
35.00
27.78
0.50
13/60
21 .67
12.97
Control
6/60
10.00
—
Ca rba ry1
10.00
48/60
80.00
77-36
1.00
41/60
68.33
64.15
0.50
40/60
66.67
62.27
0.25
31/60
51.67
45.28
0. 10
20/60
33.33
24.52
Control
7/60
11.67
—
Ma 1 athion
10.00
52/60
86.67
85.46
7.00
48/60
80.00
78.18
4.00
35/60
58.33
54.54
1.00
29/60
48.33
43.63
0.50
16/60
26.67
20.01
0.10
13/60
21.67
14.55
Control
5/60
8.33
—
Note: Each
15 mature female
test concentration was
northern fowl mites per
replicated four
repl¡cation.
times using

Probits
Figure bO. Probit curves for all acar¡cides tested, plotted on one set of axes.

198
Table 55- LCso's and regression equations for the acaricides tested.
Acá ricide
LC50 (ppm)
Reg ression
Equation
TM
SBP 1382
3.0
Y = 11.46
+ 0.80X
BW 21 Z
2.5
Y = 13.02
+ 0.97X
SD 43775
4.4
Y = 12.70
+ 0.99X
TM
Ect1 ban
1.8
Y = 11.51
+ 0.75X
Ca rba ry1
0.4
Y = 7.90
+ 0.29X
Malath ion
1.7
Y = 9.15
X
co
-rr
0
+
Note: Acaricides were tested against the northern fowl mite.

199
The LC 5o1s of the four synthetic pyrethroid compounds were very
closely grouped (Table 55). Ectiban had the lowest LC50 at 1.8 ppm and
SD 43775 had the highest at 4.4 ppm. The slopes of the synthetic pyre¬
throid curves were also closely grouped and approaching 1.0 (Table 55,
Figure 40). This indicates that the susceptibility of the mite popu¬
lations to these compounds was relatively heterogeneous. This may be
due to chlorinated hydrocarbon cross-resistance which has been reported
for other acari (Nolan et al., 1977).
The LC5o of malathion at 1.7 ppm was only slightly lower than the
LC5o of Ectiban, but the LC50 of Sevin at 0.4 ppm was lower than the LC50
of SD 43775 by a factor of 10. The slopes of the malathion and Sevin
curves were less than 0.5, thus the curves were flatter than the syn¬
thetic pyrethroid curves (Table 55, Figure 40). This indicates that
the susceptibility of the mite populations to these two compounds was
more heterogeneous than it was for the synthetic pyrethroids, and that
the mite populations are in the process of becoming more resistant to
Sevin and malathion (Hoskins and Gordon, 1956). Northern fowl mite
resistance to malathion was recorded by Hall et al. (1978).
Although Sevin had the lowest LC50, it had the highest LC90 of all
compounds tested (Figure 40). In order to produce mortalities of up to
100% in the field, the concentration of Sevin would have to be much
greater than the LC100 concentrations of malathion or the synthetic
pyrethroids.
Control of Endemic Florida Strains of Northern Fowl Mites with Carbary! ,
Malathion, and Ravap
Data were collected from one hen in each of the first 120 cages on
the north side of the four Ca1 ifornia-sty1e houses at the tilling site.

200
All three hens in each cage were examined initially, and the one having
the highest mite population was selected as the test animal. For identi
fication purposes, test animals were marked on the wing feathers with a
water-proof black magic marker. A total of 480 hens was used in the
experiment.
The formulations and mixing instructions, test concentrations and
application rates for the acaricides tested are shown in Table 56. The
treatment schedule is shown in Table 57- Malathion in treatment 1 was
incorrectly mixed and applied before the error was discovered, as
frequently occurs on commercial poultry operations. A second malathion
treatment was added with malathion mixed at the proper concentration.
Both treatments were maintained throughout the experiment.
On the day before the initial application of acaricides, a mite
precount was made. Two weeks after making the precount, a second mite
count was made. The following day, the second treatment was applied.
Two weeks after making the second mite count, a third count was made,
followed by a third treatment the following day. Two weeks after the
third mite count was made, a fourth and final count was made.
Resu Its. Weekly mite population means and converted population
means are shown for each treatment in Table 58. A plot of mite popula¬
tion means per hen by treatment is shown in Figure 4l .
There were significant differences between precount mite population
means in the treatment groups (Table 58). This was unimportant, however
because the precount mite mean for the control group was in between the
highest and lowest precount mite means, and because the final mite
counts for all groups were significantly lower than the precount of the
control group.

Table 56. The formulations, mixing procedures, test concentrations, and application rates for
acaricides tested at the tilling site for control of northern fowl mites.
Acar icide
Formu1 a tion
Volume of
Pest icide
Volume of
H20 (ml)
Test
Concentra tion
(%)
Appl¡cat ion Rate
Malathion
56.IS EC
30 m 1 3
38AO
0. bk
38AO ml/60 to 80 birds*3
Carba ry1
50.0% WP
22.7 g
38^0
0.30
38AO ml/60 to 80 birds
D TM
Ravap
28.7% EC
¿18 ml
38^0
0.36
38^0 ml/60 to 80 birds
Malathion used in week 0 of treatment 1 was 6 times (6X) this volume.
Even though each replication contained 20 test birds, 60 birds were actually present and taken
into account during acaricide application.

Table 57- Treatment schedule of acarlcldes tested at the tilling site for northern fowl mite control.
Treatment No.
Week 1
Week 3
Week 5
1
Halathlon (6X)
Malath ion (IX)
Malathlon (IX)
2
Control
Con tro 1
Control
3
Carbary 1
Carbary 1
Carba ry1
k
Carbary 1
Ca rba ry1
Carbary 1
5
„ TM
Ravap
Ravap
Ravap
6
Malathlon (IX)
Malath ion (IX)
Malathlon (1X)

Table 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.
Coun t
Number
1
2
T rea tmenta
3 4
5
6
Mi te popu1 at
ion means per hen from field counts
1 (Precount)
179.36A
116.88BCD
102.88C
18|.46A
I60.61AB
82.630
2
20.43CDE
110.78A
50.89B
28.01 CD
3.63E
34.00BC
3
19.00C
82.94A
8.23CD
34.20B
0.46D
15 - 81 CD
4
6.81 B
58.69A
3.42B
2.22B
3.13B
0.00B
Converted population means per hen^
1 (Precount)
1458.40
950.19
836.37
1478.64
1305-78
671.74
<■»
4
166.06
900.60
413.71
227.75
29.55
276.42
3
154.47
674.29
66.89
278.07
3.76
128.48
4
55.39
477.13
27.79
18.01
25.41
0.00
treatment 1 = ma 1 a th i on (6X)-ma 1 a th ion (1 X)-ma 1 a th i on (1X) .
Treatment 2 = control-control-control.
Treatment 3 = carbary1-carbary1-carbary1.
Treatment 4 = carbary1-maI athion(1X)-carbary1.
Treatment 5 = Ravap-Ravap-Ravap.
Treatment 6 = malathion(1X)-ma1athion(IX)-malathion(1X).
^Converted population mean = mite population mean from field count X 8.13.
Note: Mite population means per hen from field counts in the same row having unlike letters are
significantly different (p < 0.05) using the method of J. V/. Tukey.

11TE HEANS
Count
Figure 41 .
TM
Hite population means from hens treated with malathion, carbaryl, and Ravap at
the tilling site.

205
Two weeks after the initial treatments were applied, the mite popu¬
lation means for all treatment groups were significantly lower than the
mite population mean of the control group (Table 58, Figure Hi). Signi¬
ficant differences also occurred between the means of treatment groups.
The Ravap treatment group mite mean of 3.63 mites per bird was numeri¬
cally lower than the mite means of the other groups. The next highest
group was the 6X malathion group with a count of 20.43 mites per bird.
The remaining groups were the carbary1-ma1athion(1X)-carbary1, malathion
(1X)-malathion(lX)-ma1athion(1X), and carbary!-carbary1-carbaryl groups.
The mite mean of the control was 110.78 mites per bird.
Two weeks after the second treatments were applied, the mite means
for all treatment groups were still significantly lower than the mite
mean of the control group (Table 58, Figure 4l). Significant differences
also occurred between means of treatment groups. The Ravap treatment
group mite mean of 0.46 mites per bird was numerically lower than the
mite means of the other groups. The next highest counts were in the
carbary1-carbaryl-carbaryl group, followed by the two malathion groups
and the carbaryl-ma1 athion(1X)-carbary1 group. The mite mean of the
control group was 82.94 mites per bird.
At the time of the last mite count, mite population means of all
treatment groups were significantly lower than the mite population mean
of the control group (Table 58, Figure 4l) and no significant differences
occurred between treatment groups. The ma1athion(1X)-ma1 athion(1X)-
malathion(1X) group mite population mean was reduced to zero. The next
highest group was the carbary1-ma1athion(1X)-carbary1 group followed by
the Ravap-Ravap-Ravap, carbaryl-carbaryl-carbaryl, and malathion(6X)-

206
malath ion(1X)-ma]athion (1X) groups. The control group had a mean mite
count of 58.69 mites per bird.
All treatments significantly reduced mite populations in 6 weeks,
but Ravap reduced mite populations more dramatically than the other
acaricides and in a shorter period of time.
Efficacy of Two Synthetic Pyrethroid Compounds Against Northern Fowl
Hites on Laying Hens in Floor Pens
Seventy-five 41-week-old White Leghorn laying hens at the University
of Florida Poultry Science Research Farm in Chipley, Florida, were
randomly divided into three groups of 25 hens each. Groups were housed
in pens measuring 3-85 by 3-65 m with wood shavings as floor litter.
Natural light was supplemented with morning and evening incandescent
lights so hens were exposed to 15 hr of light per day. A 16.9% protein
laying mash and water were provided free choice. Eggs were collected
daily by pen.
The synthetic pyrethroid test materials, mixing procedures, and
application rates are shown in Table 59- Immediately following a pre¬
treatment mite count on each hen, treatments were applied. A post¬
treatment mite count was conducted 36 days later. Prior to treatment,
birds were leg-banded to identify them during the pre- and post-treatment
m i te counts.
ResuIts. Pre- and post-treatment field-estimated and converted
mite population counts and treatment means for each treatment are shown
in Table 60. Pre- and post-treatment field-estimated mite population
means are represented graphically in Figure 42.
There were no significant differences between pretreatment mite
population means (Figure 42), and the mite populations on the control

207
Table 59- Formulations, mixing procedures, and application nates for
synthetic pyrethroids applied to floor birds in Chipley,
FI ., for northern fowl mite control.
Synthetic
Volume of
Volume of
Final
Application
Pyrethroid
Pesticide(ml)
Water (1)
Solution(%) Rate
BW 21Z (42.5% EC)
4.5
3.84
0.05
3.84 1/100 birds
SD 43775 (42.5% EC)
9.0
3.84
0.10
3.84 1/100 birds

208
Table 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, FI., for northern fowl mite control.
Pretreatment Counts
Post-treatment Counts
Observation
Field
Estimates
Converted3
Field
Estimates
Converted
Treatment 1
(BW 21Z)
1
999
3121.87
0
0
2
500
4065.00
0
0
3
100
813.00
0
0
4
100
813.00
0
0
5
25
203-25
0
0
6
50
406.50
0
0
7
100
813.00
0
0
8
75
609.75
0
0
9
100
813.00
0
0
10
100
813.00
0
0
11
100
313.00
0
0
12
250
2032.50
0
0
13
100
813.00
0
0
14
1 10
894.30
0
0
15
50
406.50
0
0
16
130
1056.90
0
0
17
100
813.00
0
0
18
O
o
2439.00
0
0
19
65
528.45
0
0
20
55
447.15
0
0
21
175
1422.75
0
0
22
100
813.00
0
0
23
110
894.30
0
0
24
150
1219.50
0
0
25
100
813.00
0
0
Hite Mean
161.76
I3i5.ll
0
0

209
Table 60. Continued.
Qbservation
Pretreatment Counts
Post-treatment Counts
Field
Estimates
Converted
Field
Estimates
Converted
Treatment 2 (SD 43775)
1
25
203-25
0
0
2
500
4065.00
0
0
3
100
813-00
0
0
4
30
243.90
0
0
5
75
609.75
0
0
6
120
975.60
0
0
7
120
975.60
0
0
8
200
1626.00
0
0
9
100
813.00
0
0
10
100
813.00
0
0
11
50
406.50
0
0
12
150
1219.50
0
0
13
75
609.75
0
0
14
150
1219.50
0
0
15
200
1626.00
0
0
16
95
772.35
0
0
17
150
1219.50
0
0
18
30
243-90
0
0
19
65
528.45
0
0
20
150
1219.50
0
0
21
50
406.50
0
0
22
50
406.50
0
0
23
150
1219.50
0
0
24
75
609.75
0
0
25
35
284.55
0
0
Mite Mean
113.0
918.69
0
0

210
Table 60. Continued.
Pretreatment Counts
Post-treatment Counts
Observation
Field
Estimates
Converted
Field
Estimates
Converted
Treatment 3
(control)
1
50
406.50
125
1016.25
2
75
609.75
175
1422.75
3
35
284.55
75
609.75
4
35
284.55
90
731.70
5
110
891.00
75
609.75
6
25
203.25
65
528.45
7
65
528.45
300
2439-00
8
50
406.50
95
772.35
9
55
447.15
175
1422.75
10
25
203.25
95
772.35
11
56
455.28
185
1504.05
12
125
1016.25
300
2439.00
13
100
813-00
150
1219.50
14
180
1463.40
95
772.35
15
100
313.00
275
2235.75
16
75
609.75
255
2073.15
17
300
2439.00
250
2032.50
18
50
406.50
90
731.70
19
300
2439.00
375
3048.75
20
75
609.75
275
2235.75
21
110
891.00
300
2439.00
22
75
609.75
400
3252.00
23
150
1219.50
800
6504.00
24
50
406.50
100
813.00
25
200
1626.00
85
691 05
Mite Mean
98.84
803.57
208.20
1692.67
a
Converted mite count = field estimation X 8.13.

200
160
LO
<
UJ
LU
120
80
40
0
n
O
1
—r
i
o =
BW 21Z
n =
SD 43775
O =
Control
r~
Test Animal: northern fowl ml te
14
21
28
—r
O
35
Day
Figure 42. Pre- and post-treatment field-estimated mite population means from floor birds
treated with two synthetic pyrethroid compounds at Chipley, FI.
21 1

212
birds were numerically lower than the mite populations on birds in the
two treatment groups.
On day 36, there were significant differences between the mite
population means on the treated birds and the control birds (Figure 42).
Mite populations on the treated birds were reduced to zero, but the mite
populations on the control birds averaged 208.2 per bird.
The mean egg production by treatment by day is shown in Table 61.
Analysis of production data was run by day and by the 36-day production
mean. There was no significant difference in production due to treat¬
ment shown by either analysis.
The Effects of Northern Fowl Mites on Egg Production
Twelve commercial strains of White Leghorn laying hens on the
University of Florida Poultry Science Research Farm in Chipley, Florida,
were used to evaluate the effects of northern fowl mites on egg produc¬
tion. Birds were housed two to a cage in cages measuring 31 by 25 by
46 cm and were 41 weeks old when the experiment began.
Two houses were utilized for the trial. House 200 was a California-
style house 82.3 m long by 3-7 m wide with the center aisle on an east-
west axis. House 100 was 91.4 m long, 9-1 m wide, and oriented parallel
to house 200. House 100 was partially enclosed with aluminum siding and
wild-bird-proofed with 1-cm hardware cloth. All the hens in house 200
and the caged layers housed in the west end of house 100 were utilized
for the experiment. The number of hens tested ¡r. each house was the
same.
Both houses contained four rows of stair-step cages (Figure 43).
Each row was divided into twelve 24-cage blocks with 12 cages on the
upper row directly above 12 on the lower row. The 12 strains of hens

213
Table 61. Daily egg production means of birds treated with two
synthetic pyrethroids in Chipley, FI.
Day
1
T reatment
2
3
2
0.24
0.24
0.56
3
0.56
0.34
1.04
4
0.92
0.80
0.80
5
0.80
0.84
0.80
6
0.64
0.64
0.68
7
0.92
0.88
0.76
8
0.72
0.48
0.88
9
0.40
0.88
0.32
10
1 .00
0.80
0.76
11
0.76
0.76
0.64
12
0.80
0.76
0.60
13
0.76
0.52
0.76
14
0.92
0.92
0.80
15
0.52
0.52
0.60
16
0.80
0.80
0.60
17
0.60
0.44
0.56
18
0.84
0.64
0.76
19
0.84
0.56
0.92
20
0.60
0.48
0.44
21
0.80
0.76
0.80
22
0.92
0.80
0.76
23
0.56
0.56
0.36
24
0.72
0.68
0.52
25
0.68
0.44
0.56
26
0.60
0.84
0.80
27
1.20
0.84
1 .00
28
0.64
0.68
0.52
29
0.68
0.56
0.72

214
Table 61. Continued.
Day
1
T reatment
2
3
30
0.80
0.56
0.72
31
0.64
0.40
0.58
32
0.88
0.84
0.72
33
0.68
0.64
0.64
34
0.72
0.24
0.36
35
0.96
0.48
0.56
36
0.52
0.60
0.40
Note: Treatment 1 = BW 21Z at 0.05?
Treatment 2 = SD 43775 at 0.10?
Treatment 3 = Control
Treatments were applied to floor-housed birds.

House 100 (birds in west end onl.y)
N
Treated
I 1 Control
Figure Houses 100 and 200 showing locations of strain replications and treatment areas.
215

216
were randomized throughout the houses so that each row of cages housed
one 24-cage block of each strain. Blocks were divided in half. Hens
in one half received a diet with a varying protein level and hens in
the other half received a diet with a fixed protein level.
All hens on the lower rows of cages in house 200 were examined for
northern fowl mites. One bird in each cage was marked on the wing with
a black water-proof magic marker so both birds could be identified indi¬
vidually during each mite examination. Mite populations were counted
on the day of but prior to the first acaricide treatment and again 36
days later (Table 62). Mites on birds in house 100 were not monitored.
Egg production records were maintained in both houses. Eggs were
collected daily and production was calculated on a hen-day basis for
both feed protein levels within each block. Because of results from
previous trials, Ravap was the acaricide of choice. The test concen¬
tration of 0.36% was mixed by adding 48 ml of Ravap 28.7% EC to 3.8 1
of water. The application rate was 3.84 1 per 60 to 80 birds. Ravap
was applied on days 1 and 29 (Table 62) to hens in rows 2 and 3 in
both houses (Figure 43).
Morning and evening artificial lights supplemented natural light
and provided hens with a total of 15 hr of light per day. Water and
feed were offered ad libitum.
TM
A fly control system, manufactured by Chem-amatic in Bellview,
Florida, had been installed under the cages in house 200. When an
automatic timer activated the system, a 0.033% pyrethrin solution was
dispensed through spray heads located at fixed intervals throughout
the house. The height of the spray heads was just above the level of
the walks.

Table 62. Data collection and treatment application schedule for the
Ravapâ„¢ northern fowl mite trial in Chipley, FI.
3 days prior to pre¬
treatment mite count
Began collection of egg production
data on a daily basis.
Day 1
Mite pretreatment count completed;
Ravap was applied.
Day 29
Second Ravap application.
Day 36
Post-treatment mite count completed.
Day 45
Terminated collection of daily egg
production data.

218
ResuIts. With the exceptions of strains 8 and 9, Ravap reduced
mite populations to zero with two applications, as shown in Table 63.
Mite numbers were reduced significantly (p < 0.0001) in all cases.
Although there were no significant differences between mite popu¬
lation means of treatment groups prior to treatment, significant
differences between strains did occur (Table 64). This was due to the
wide variation in mite numbers found on strain replications in the four
quarters of house 200. Table 65 shows a significant difference between
post-treatment group mite means. Mote that the mite populations in the
untreated group had doubled since the pretreatment count. Also shown is
the ranking of mite means by strain. This is essentially a ranking of
means from the untreated groups, since the mite populations in the
treated groups had been virtually eliminated. Means shown in Table 65
were reduced by a factor of 2 since both treatments were considered in
their computation.
In Table 66, pre- and post-treatment mite counts in the control
group were added together and the means computed. This table gives some
indication of strain resistance to mite populations. The strains, as
ranked here, either maintained high mite populations throughout the
trial or had rapid increases in population numbers during the trial.
Upon our arrival in Chipley, Florida, to begin the pretreatment
mite count, it was noted that the pyrethrin fly control system previously
mentioned was in partial operation on the north side of house 200.
Since one group from each treatment was on the north side of the house,
the house was divided into east and west halves and both halves were
analyzed separately. Tables 67 and 68 show the pretreatment mite counts
by treatment and strain for the west and east ends respectively of house

Table 63. Pre- and post-treatment mite population means by treatment and strain from caged-layer trial
at ChipIey, FI.
Pretreatment
Post
-treatment
Strain
Trt Ia
i-xb
Trt 2
2-X
Trt 1
1 -X
Trt 2
2-X
1
10.ii
84.6
1541.7
12534.0
0.0
0.0
742.0
6032.5
2
37-7
306.5
1954.2
15887.6
0.0
0.0
1731.8
14079-5
3
66.0
536.6
1279.7
10404.0
0.0
0.0
1891.7
15379.5
4
202.2
1643.9
414.9
3373.1
0.0
0.0
283.7
2306.5
5
500.0
4065.0
729.4
5930.0
0.0
0.0
728.1
5919.5
6
3714.6
30199.7
278.7
2265.8
0.0
0.0
645.2
5245.5
7
4151.0
33747.6
314.9
2560.1
0.0
0.0
1289.9
10486.9
8
2530.3
20571-3
562.5
4573.1
55.5
451.2
1063.5
8646.3
9
1447.9
11771.4
170.8
1388.6
0.5
4.1
4183.4
34011.0
10
2234.0
18162.4
59.6
489.5
0.0
0.0
3464.4
28165.6
1 1
839.6
6825.9
140.2
1139.8
0.0
0.0
1790.8
14559.2
12
351 .1
2854.4
3666.7
29810.3
0.0
0.0
5896.9
47941.8
3 Treatment
T reatment
1 = Ravap
2 = Control
Mite population means
converted
to actual
mean mite
number by
process
dése ribed
in text
(actual mean = mite population mean X 8.13).

220
Table 64. Pretreatment mite population means by strain (treatment
ignored) and by treatment group from the caged-layer trial
at Chipiey, FI.
G3CU
p r ng
I3
N
STFa:>
A
225 3.
157895
95
7
A
B
A
202o .
315739
* '15
1 2
Ü
A
3
A
20 1 A.
73 6 2
9 5
5
3
A
p
A
1 5^6.
9 ó
9
Q
A
c
A
c
1 I ió .
30 3 5 1 1
r* a
. ->â– 
1 0
3
r
3
/â– -
9 95.
93 " 5 0 0
95
o
V.
c
30 9 .
375000
96
9
c
c
77 6 .
1 óó 7
9 5
i
c
5 7 .
210526
9 5
J
£
>:> 1 A •
533333
o
/“
c
A A 3 •
634211
95
l 1
.30 => .
677 3 1 9
93
b
G - OijP ING
AN M
p T
A
tj-6.732537 57i
l
r*
A
132.092320 57\
p
a
Means with the same
letter are not significantly
d ¡ fferent
(p < .05).
k Treatment 1 = Ravap'^, Treatment 2 = Control.

221
Table 65. Post-treatment mite population means by strain (treatment
ignored) and by treatment group from the caged-layer trial
at Chipiey, FI.
GROUPI
NG
'â– 'ZA'i 3
N
STPAI
A
2^7 9 .
27 6 3 2
95
1 2
n
o
2 113.
957395
0 5
3
CJ
0
1 73 2.
l80351
94
1 0
c
92 5 .
533333
9 ó
3
c
c
27 6.
32 9 7 3 7
9 4
1 1
c
c
86 3.
535-17
°5
-*
c
0
r
63 d.
157305
95
—f
D
c
D
r
£61.
321053
9 5
D
c
0
J?0 .
93:>533
Q 5
1
n
r
r,
C
3m-.
052500
Q --N
.5
r>
0
-
3 1 v •
210 52Ó
9 5
5
1 2 0..
925652
92
u
Gr- GU 9 I NG
V SAN
M
b
A
l9 5 0. i ^2211
5^0
'¿
-.627217
¿6 9
1
(p <
Means with the same letter are not significantly different
05) •
TM
Treatment 1 = Ravap , Treatment 2 = Control.

222
Table
66. Means of the
the control
at Chip1ey,
combined pre- and post¬
group (treatment 2) from
FI .
treatment
the caged
mite counts of
-layer trial
a
GPGUPINÚ
M " A N
M
3 T- AIN
9 563. - PI 6 7
d
1 2
B
435 -. 13^50 0
*4 d
9
e
£
3
3 v 3 5 . «3 '7 5 0 0
A. 2}
->
Q
B
P
3523.-36190
¿-7
l 0
c
d
r
B 0
31’ 1.35—lo7
4. Q.
3
c
0
c
E D
2.2 8 3 • o 4 5 B 2 3
43
1
5 0
1934. 239 180
4 6
1 1
c
162c.04iBcT
43
O
E
l 6 J 4 • ''37234
- 7
7
1 45 7.901 5A7
A CO
)
92 3. • 3c 17 0
¿l 7
6
-
693.617021
a?
—
Means with the same letter are not significantly different
(p < .05).

:'i n n n r. n o n n
223
Table 67. Transformed pretreatment mite population means by strain
(treatment ignored) and by treatment group for the west
end of house 200.
GKC'J=>i\.;3
M E A ' 1 a
rr t A T \
O 1 r-. A i .
M
42.S24778
4 7
6
30.6>0520
>c
7
P
r\
26.473322
4 7
1 0
3
r-
22.04134Q
4 S
9
0
0
p
C
2 1 .2102 = 4
4S
â– 5
3
0
3
0
2 C.ol 0 2 0 0
4 7
12
D
0
17.317422
4 =
27
F
C
F
D
12.237713
4
?
=
= .4302 1 4
4 7
1 1
cr
c . 2 i 4 7 J 0
4.5
1
3.622640
4 6
4
2.233420
4 3
5
c^ru^: n MEAN
N
t? r b
A
2 1. 5 1 1 3 C 4
2 A ó
1
5
1 5.3 733*75
2'U
2
(P <
Means with the same letter are not significantly different
05).
TM
Treatment 1 = Ravap , Treatment 2 = Control.
Note: Data was transformed using / N+l

224
Table 68. Transformed pretreatment mite population means by strain
(treatment ignored) and by treatment group for the east
end of house 200.
c u â– 
â–  I i j
7 c A N a
! I 3 A I
A
.1 A
•
612115
-4 6
q
A
A
3 3
2 £ 4 7 7 9
.a
•4 CJ
T P
A
A
2 7
•
640553
7
-3
-A
c:
A
C
*> wt
•
4 7 1 5 4.3
•; -
3
i *
A
c
jp
A
c
o c
•
0 72 7d 1
4 A
2
c
s~
\_
7 1
•
125222
45
1
n
c
1 9
a
3 7 95 34
4 3
o
Jj
c
â– 5
c
l c
a
754754
4 7
1 0
on
c
3
c
i t)
•
OCÓC3D
4
11
0
c
P
c
1 5
•
267752
4 c
â– p
23
c
q
c
1 4
772051
4 7
4
r~
v_
c
1 2
»
5451 36 1
4 7
q
37 A
A
A
22.222756 2 - 2
21.7-0110 267
b
TRT
1
a
Means with
the same letter are not significantly
d i fferent
(p < .05).
b __
T reatment
TM
1 = Ravap , Treatment 2 = Control.
Note: Data
was transformed using / N+l

225
200. The mite populations in the control group in the west end of the
house were significantly lower than those of the treated group. This
is reflected in the differences in the mean rankings by strain between
the east and west ends of the house (Tables 67 and 68). The northwest
section, where the pyrethrin spray system was partially operating, was
where the control birds were located. Tables 69 and 70 show the post¬
treatment mite counts by treatment and strain for the west and east
ends respectively of house 200. Since the mite means for the control
groups in both ends of the house were almost numerically equal it was
discounted that the pyrethrin spray system had any effect on the mite
popu1 a tions.
Egg production means by treatment and strain for house 100, house
200, and houses 100 and 200 combined, are shown in Tables 71, 72, and
73 respectively. There were no significant differences in egg produc¬
tion due to treatment when strain differences were ignored. Mean
rankings by strain were very similar in all three tables.
Table 74 gives the egg production means by week, with and without
treatment being taken into consideration. Weeks 6 and 7 were signifi¬
cantly different from each other and from weeks 1 through 5, but there
were never any significant differences between treatments at any point.
There were no differences in egg production due to feed protein level
and results are not reported.
A plot of weekly egg production means by treatment is shown in
Figure 44. A nonsignificant drop (ca. !%) in egg production occurred
3 days after both applications of Ravap (Table 74, Figure 44), possibly
due to treatment stress. In each esse, birds recovered and within 7
davs began to lay at a numerically higher rate than the control birds.

226
Table 69. Transformed post-treatment mite population means by strain
(treatment ignored) and by treatment group for the west
end of house 200.
gp cup i r ¿0
a
y l \
N
3T 5 A I
A
3 4.7 4 ? 4 1 5
4 7
9
\
b\
A
2 2.4 2 l 9 4 5
. 4 ?
1 2
.A
■¿a .52 13 01
4 7
1 0
T.
1 J • T6212c
4 7
L l
3
1 5 . c 5 "5 5 21
4 5
3
2
15.585522
4 s
3
1 4 . 645789
- 2
7
C
1?.71c3:3
4 6
1
z
1 3.6275 G5
4 3
O
c
C E
10 .520 330
4 7
Ó
C 3
1 231 445
4 ¿
5
C
5.50 7 10 4
4 '5
4
b
I NG
MEAN
T9 T
A
23.912224
2 :» 4
C-
3
1.337101
2 34
1
Means with the same letter are not significantly different
(p < .05).
b TM
Treatment 1 = Ravap , Treatment 2 = Control.
Note: Data was transformed using / N+l

227
Table 70. Transformed post-treatment mite population means by strain
(treatment ignored) and by treatment group for the east
end of
house 200.
a
»P Gl j . I ' • vj
•V £ A N
N
5 T r- 4 I 'J
A
o7.DEB7il
.x 3
1 1
24.10=411
n 5
5
P
2 4.133737
/r -3
v>
p
22.041772
4 6
2
n
c
3
2 C . 932322
4 7
1 0
c
r
5
0
16.552755
4 7
1 l
c
D
P
0
14.237567
-7
3
c
;r
0
r
:r
£
15.330753
4 7
7
p
3
p
3
1 1.442 1 03
4 5
•r-
0
D
11 ,19 10 15
4 a
5
E
5.337554
4 c
i
5
7.953779
47
4
â– o;--ooJ IN<3
A
ó
VEAN N r 3 7
34.350701 ?5o
1 . 0 G 0 3 0 C 2 15 1
3 Means with
the same letter are not significantly different
(p < .05).
T reatment
Ravap
TM
Treatment 2 = Control.
Note: Data was transformed using / N+l

228
Table 71. Egg production means by strain (treatment ignored) and by
treatment group for house 100.
GROUP I MG
PEAN 3
M
STRAH
A
B 4.2D 1454
39 2
1 0
A
A
33.422092
392
6
B
81.148036
39 2
7
a
c
a
SO. 832332
392
4
c
3
r
a
3 0.47 7321
39 2
3
c
c
0
7 5.974 031
3 92
o
1—
D
E
D
77.938139
302
5
h_
E
F
76.110230
392
9
F
G
F
74.648240
392
1 2
G
G
7 3. 5 3540 3
3 92
1 1
H
7 1 .S993~'2
39 2
i
I
6 3. 3157")-*
3-* 2
3
b
GROUPiMG
MEAN
ft
TRT
A
7 7.9916 o2
2 35 2
O
A
A
77.3 08373
2 35 2
1
Means with the same letter are not
(p < .05).
k Treatment 1 = Ravapâ„¢, Treatment 2
significant 1y
- Control.
different

229
Table 72. Egg production means by strain (treatment ignored) and by
treatment group for house 200.
a
F C;rr’
r r • j
VEAN
N
5 T K A Í n
A
33.333322
252
1 C
A
A
¿3.1 ¿-9 c 1 7
3 92
6
A
F
A
31.579239
7A 21
3
^2
r~
30.625255
3 92
7
C
73.937500
39 2
2
D
7c.ic6t?ó'i
352
9
5
7 fi . j 0 S 5 óo
352
5
rr
r\
E
D
7c ~ 1 E 3 3 2
3 92
4
E
F
74.469032
392
l 2
p
E
7 4. C6 71 17
72 C P
—.7-4—
1 1
F
7 1.422602
392
1
G
62.4224 IS
352
O
u
b
GPOUP I NG
MEAN N TPT
A
77.062953 2352 1
A
A
76.667721 2352 2
3 Means with the same
letter are not significantly different
(p < .05).
k Treatment 1 = Ravap"^, Treatment 2 = Control.

230
Table 73- Egg production means by strain (treatment ignored) and by
treatment group for houses 100 and 200 combined.
â– o K UU <1 I N 'o
a
i' i L. r-y,' i
5 TP \ T '
/A
33.76 7533
754
1 0
A
A
33.290355
7 3 -+
6
0
3 1.0 2 3 26 5
7 34
~J
t»
3
30.836035
73 +
7
c
73.930765
7 3 4.
O
c
c
73.039032
7 A :t
%
D
7 6 » 6 7.3 8 7 3
73 4
3
D
D
76.403597
7 3'4
9
=
74.552771
7 3
1 2
e
E
7 3.3262 >3
73 +
1 1
F
71.511237
7 3 4
O
ó 3•o2 2 5 64
734
s
r 1
b
T7 r
G K Ü ’J/-7 I V To
A
77.4 2'1'^ 72
4-7 J 3
—>
A
A
77,153443
4 7 j 4-
(p <
Means with the same letter are not significantly different
05).
TM
Treatment 1 = Ravap , Treatment 2 = Control.

Table
(p < •
231
74. Egg production means by week (treatment ignored) and by
treatment group
in the caged-layer trial at
Chipley.
: r i J I \ i / '
U t A • J ' J
"i f. a : 4 a
9
V/ p' r‘
A
79. 7 7 0^0 1
1 2 -t i
â– >
A
A
79 .o03ü53
13+4
Ó
A
•
A
79.53356 1
13.4
A
A
A
79.512312
1 5 +4
A
A
79 . A 3ó° 7 o
1 3 4 4
i
.3
77.614359
13 44
r.
C
65 • o i 7" 4 Q A
1 3 4 3
7
LY
PR OCUCTI ON
M ; A i'iS
BY TP-AT
r^S
•VFEK
_ „ b
ME 5 5
1
1
A.
1
7 3.4 961
2
1
O
r_
79.5-19
3
9
l
79.7445
A
2
£_
7 3.7 6^4
5
3
1
7 9.3139
o
p
7 3.7 3-7 ;•
~7
4
l
79.4197
' J
7 7. 3 909
9
a.
Z)
1
73.711°
1 0
* 5
79.2123
1 1
o
i
7. 6 A 3 0
1 2
A
'»
C_
7'1’. 1 690
l 3
7
1
6 5.9020
1 4
7
2
¿4. 3=17
Means with the same letter are not
05).
TM
Treatment 1 = Ravap , Treatment 2
significan11 y
= Control.
different

Production Means
Figure Weekly egg production means by treatment from the caged-layer trial at
Chipley, FI.
K>
OJ
ho

233
The decrease in egg production beginning in week 5 was attributed to the
beginning of an outbreak of fatty liver syndrome (R. H. Harms, personal
communication). Patent symptoms did not appear until several weeks
after the experiment ended.
»
Analysis of individual strains by a t-test revealed a significant
difference in production in one strain due to treatment (Table 75). The
treated birds in strain 4 were producing eggs at a level slightly above
the average production level of the 12 strains combined. The production
level of the treated strain 4 hens represented a 3.67% increase in pro¬
duction (p < 0.0137) over the untreated strain 4 hens. As can be seen
in Table 77, and other tables previously mentioned, strain 4 hens main¬
tained low mite populations throughout the trial. Strain 4 was also
one of the few strains to exhibit a natural decrease in mite population
at the end of the trial (Table 77).
In an effort to correlate production data with effects caused by
the previously mentioned pyrethrin sprayer, each quarter of house 200
was analyzed as a different treatment, i.e. treatments 1 through 4.
Treatment means (treatments 1 through 4) shown in Table 76 were not
significantly different and again, any effects of the pyrethrin sprayer
were discounted.
In an attempt to show a correlation between mite numbers and egg
production, a plot of egg production means versus mite means by strain
was made (Figure 45). According to this graph, there appears to be no
correlation. Birds with mite populations of ca. 640 or ca. 2320 mites
per bird were both laying at the 81 percentile.
However, it appears that it is not mite numbers alone that affect
production, but a combination of mite numbers and time (Table 77).

234
Table 75-
T-test of egg production treatment means
caged-layer trial at Chipley, FI.
by strain from the
Strain
T reatment3
Mean {%)
Ab
Prob < T {%)
1
1
70.46
-I.92
82.53
2
72.38
2
1
78.32
-1.34
63-43
2
79-66
3
1
81.80
+0.44
24.09
2
81.36
4
1
77-05
+3-67
98.63
2
73-38
5
1
76.42
+2.03
84.34
2
74.39
6
1
83-03
-0.26
14.05
2
83-29
7
1
80.47
-0.31
17-01
2
80.78
8
1
69-15
+ 1.44
70.15
2
67-71
3
1
76.84
-0.09
4.79
2
76.93
10
1
83-52
+0.37
20.62
2
83.15
11
1
73-20
-1-73
76.80
2
74.93
12
1
74.48
+0.02
1.30
2
74.46
x =
= 76.97
3 Treatment 1 = Ravap"^,
Treatment i
’ = Control.
Mean difference between treated and untreated birds.

235
Table 76. Egg production means when each quarter of house 200 was
analyzed as a separate treatment.
ürüUPINo
N TRTb
A
7 7 . ó 6 o a 0 2
11c]
"7
A
A
76.391932
1 1 70
4
A
*
A
76.c,il5l4
1 1 76
o
A
A
7ó .432A?7
119 1
l
Means with the same letter are not significantly different
(p < .05).
b Treatment 1 SW = Ravap^, Treatment 2 NW = Control + pyrethrin,
Treatment 3 NE = RavapTM + pyrethrin, Treatment 4 SE = Control.

Table 77- F.gg production means, pre- and post-treatment mite population means, and the change in mite
population numbers on untreated hens from the caged-layer trial at Chipley, FI.
Strain
Pretreatment
Mite Mean
Post-treatment
Mite Mean
Change in Mite
Population During Trial
Overa 11
Egg Production
Mean {%)
6
278.7
645.2
t
2. OX
83.29
iob
59-6
3464.4
t
X
0
0
LTV
83.15
2 a, b
1279.7
1891-7
T
1 - 5X
81 .36
7
31^.9
1289.9
t
4. OX
80.78
2a»b
195*t.2
1731.8
f
0.9X
79.66
9b
170.8
4183.4
f
25.ox
76.93°
llb
140.2
1790.8
'1'
12. OX
74.93
I2a’b
3666.7
5896.9
t
1.5X
74.46
5a
729.4
728. 1
No
• Change
74.39
A
414.9
283.7
+
0.7X
73.38
Ia
1541.7
742.0
+
0.5X
72.38
8a
562.5
1063.5
t
2. OX
67.71
Six strains with the highest pretreatment mite counts.
b Six strains with the highest post-treatment mite counts.
Q
Mean egg production: 76.87%-
Note: Arrows (f,f) indicate an increase or decrease in egg production.

CO
LU
21
O
O
23
Q
O
tic
CL.
cr>
LU
89
80
76
72
68
I
i 1 - 1 - r r
—1 ,—
i
t 1 r
1 r r
1 1
-
10-0
6-0
-
-
3 -0
-
7 - 0
-
9-0
2-0
-
it - 0 5-0
12-0
n~o
1 - 0
» —
8-0
1
1 1 1 11
1 1
1
1 1 1
1 1 1
1 1
0
320 m
960
1280
1600
1920 2290
2560
Mite Means by Strain
Figure 1|5. Plot of egg production means vs. precount mite means by strain from caged-layer
trial at Chipley, FI. (production means have not been adjusted to reflect strain
d i f ferene.es) .
N>
IjO

238
Four out of the six strains with the highest pretreatment mite counts
were laying below the average production level at the end of the trial.
All six strains showed little or no change in mite populations during
the trial. Of the remaining six strains, four of which were laying
above the average, only one strain, strain 4, had a mite population
change of less than +2X. Only two strains, strains 2 and 3, produced
above the production mean and maintained mite populations ranking among
the six highest in the pre- and post-treatment counts.
No matter what conclusions may be drawn from Table 77, it is inter¬
esting to note that the treated counterparts of the strains in Table 77,
all with different pretreatment mite populations, had their populations
reduced to zero or close to it. Yet, only one strain had a significant
increase in production due to treatment. A longer trial including
initial development of mite populations is necessary to more completely
investigate the factors involved in this phenomenon.

DISCUSSION
The Value of Rotovation as a Method of Manure Management
After three years of experience dealing with tilling and manure
management, I have made the following observations concerning rotovation
as a manure management tool .
The Use of Tilling for Drying Manure
Manure may stay dry without tilling. Tilling manure does not
necessarily guarantee that manure will be dry (McKeen and Rooney, 1976),
and abstention from tilling does not necessarily guarantee that manure
will be wet. During the 2 years that tilling experiments were conducted,
manure at the tilling site was very wet. These normally unfavorable
conditions were very suitable for our experimental purposes. After
tilling experiments had been terminated, a new flock of chickens was
housed and the manure the following year was the driest it had been in
over 3 years.
The wet conditions during the first 2 years were attributed to two
main factors. First, the summers were unseasonably hot. On one particu¬
lar day, 50 chickens died as a result of the heat. Second, in an
effort to cool themselves, chickens drank large amounts of water which
caused droppings to be semiliquid throughout the spring, summer, and
fal 1 .
Rotovation as a Management Tool
Rotovation is not a substitute for good management practices. in¬
stead, it is an additional management practice which can be used to dry
239

manure and reduce fly levels. Poultrymen who think that rotovatlon can
be used as a shortcut to make up for lack of continuous sound management
will soon discover that fly and manure problems will continue to exist
even after the purchase of a tiller.
Tilling once or twice a month will not correct problems caused by
leaking waterers or water seepage. In fact, tilling on an infrequent
and random basis can be worse than not tilling at all. If manure is
fairly dry and crusted over, periodic tilling will only expose wet
areas beneath the crust and encourage fly breeding. This is particular¬
ly true during the months of July through September.
In the experiments performed in this study, most tilling was done
in manure that was wetter than preferred. With sufficient time and
effort, wet manure can be tilled enough to improve its consistency and
make removal possible. This is not the purpose for which the tiller
was designed.
If the decision is made to utilize rotovation as a manure drying
technique, it must be implemented at the right time to ensure success.
Implementation of a Tilling Program
The time to begin a tilling program is in between flocks. As soon
as a flock has been removed from the farm, work should begin to properly
prepare the manure collection areas under the cages for a tilling pro¬
gram. When the old manure is cleaned out, a layer of manure should be
left under the cages at a level that the tiller's tines can penetrate.
This is also necessary to maintain a balance in biological control
agents (Peck and Andersen, 1970). Tilling should begin immediately to
dry this manure base as much as possible prior to the arrival of the new

241
flock. (Poultry farms are usually vacant 4 to 6 weeks between flocks
and flocks are changed every 15 to 18 months.) When the new flock has
arrived, the tilling program will have already begun and the manure
drying process can begin on the first day.
If a poultryman waits until a flock has been housed before deciding
to till, he has missed the opportunity to prepare dry beds in which to
stir the manure. If he waits longer before deciding, he also has to
contend with the layer of wet manure being deposited during every day he
has waited. Extra time and effort will be necessary to bring manure
moisture levels below those preferred by house flies.
In trials performed at the tilling site, tilling always began after
a flock was housed. Manure was tilled a maximum of twice a day with a
30 min time interval between til lings. The moisture content of very
wet manure was reduced to almost acceptable levels within 3 weeks.
Tilling more than twice a day with longer time intervals should help
shorten the drying period, but operation and labor costs are increased.
Tilling Frequency
Tilling frequency depends on the condition of the manure. Powdery
dry manure is routinely tilled once a day by poultrymen in the Tampa
area of Florida (C. F. Hinton, personal communication). Wet manure
should be tilled two or more times daily depending on the individual
situation. Except with semi liquid manure, progress can be seen within
1 week if manure is tilled frequently enough.
Tilling during rainy weather is not recommended especially if
manure is not dry and dusty. Once manure is dry enough to be dusty,
wetting it again is very difficult. On occasions when rain blew into

242
houses at the tilling site which contained manure in the state of dry¬
ness mentioned above, manure remained essentially unaffected. Tilling
was resumed the following day if the weather cleared.
As wet manure loses its moisture content, it breaks into particles.
The particle size will be quite large initially, but wi 1 1 continue to
decrease as manure dries. When manure becomes powdery dry, particles
essentially cease to exist and manure has a texture similar to coarse
sand.
Particle formation hastens the drying process by increasing the
manure surface area; it also allows parasitic wasps to more thoroughly
search the manure for pupal hosts (Hinton, 1977)- Particle formation
provides the poultryman with visible evidence that the manure is drying.
When to Till
Tilling should be done during the heat of the day to increase the
water evaporation from freshly turned manure. However, if manure is
wet, tilling in the morning before egg collecting begins is beneficial
as well as tilling several times during the afternoon. All tilling at
the tilling site was done between 12 noon and 2 p.m. while the egg
gatherers were at lunch. This allowed the entire farm to be tilled
non-stop without interfering with the egg pickup.
Leave a Dry Base When Poultry Houses are Cleaned Out
A dry base that the tiller's tines can reach must be left each time
manure is removed from poultry houses. Laborers cleaning out poultry
houses at the tilling site always removed the dry manure base and some
of the sand beneath it. More than one tilling experiment was halted
because the tiller's tines could net touch the bottoms of the manure
collection areas after the houses were cleaned out.

243
Two factors were responsible for this problem. First, the laborers
prided themselves in doing a thorough job of removing all the visible
manure plus a little more. Second, the poultryman refused to monitor
the activities of the laborers and insist that manure be removed only
to the level that he specified. As an observer, I could only make
suggestions that often went unheeded.
Tilling Reduces Manure Volume and the Number of Clean-Outs Per Year
Poultrymen are currently cleaning out manure three times during the
15-month period that a flock is housed. However, regular use of a til¬
ler has reduced clean-outs on some north Florida poultry farms to two
per 15-mcnth period. The present cost of cleaning out a poultry house
is $3.28 per m, or $300 for a 91.^ m house. Reduction in the number
of clean-outs per flock by one third would save the owner of the tilling
site $ 1200 per year.
This savings is increased when one considers the value of the manure
as a fertilizer (Hinton, 1977). In 1976, farms in the Tampa area were
selling rotovated manure for $10 per ton and attempts were being made
to increase the interest of the citrus industry in this readily available
product.
Rotovated manure can be spread on fields or sold in bags for ferti¬
lizer without further drying. In this form, the manure is unattractive
to flies and does not burn plants or pastures (C. F. Hinton, personal
oommunicat'ion) .
Tiller-Related Costs and Returns
Initial cost of purchasing a tiller is high but operational costs
are low. Selpat sells the tiller and the modified Kuboda as a unit,

244
although the two can be purchased separately. The price in 1976 was
approximately $3000. This capital can be regained in 3 to 5 years if a
tilling program is begun and adhered to. Returns are in the forms of
reduced manure clean-out schedules (described in the^previous section),
increased value of manure as a fertilizer (also described above) and
the reduction of adult and immature fly control costs.
An average of 33.68 min were required to till all four houses at
the tilling site. Average time required to till one 91.4-m poultry
house was 8.42 min (Table 78). Time values in Table 78 also represent
time spent moving equipment and egg carts out of the path of the tiller,
so actual time spent tilling was slightly less than the time indicated.
Some time was also spent keeping up public relations with the farm
owners.
Approximately 0.25 1 of fuel and 0.14 tractor hr were required to
till one house (Table 78). All four houses at the tilling site could be
tilled once a day for 16.55 days on one tank of fuel (19-2 1).
Using the figures given in Table 78 and assuming the cost of fuel
and labor to be $0.26/1 and $3 per hr respectively, it would cost the
owner of the tilling site $640.58/year to till his four California-
style houses once a day. This translates to $0.032/bird per year, which
is a reasonable cost for fly control. This figure quickly increases,
however, if houses must be tilled more than once a day. Hinton (1977)
quotes tilling costs of $.03 to $.10 per bird as compared to $.06 to
$-08 per bird for the purchase of pesticides. His figures are for a
50,000-bird flock.

245
Table 78. Mean operator and tractor time and the amount of fuel required
to till one 91.4-m Ca1ifornia-style poultry house.
Operator Time (min)‘
20.00
8.00
10.00
6.67
7.33
6.00
Odometer
Fue 1
5.67
8.00
Reading(hr)
Uti1ized (1)C
7.75
7-76
Start
71.90
19.20(fu11)
10.00
7.00
Finish
80.50
0.96
7.00
7.00
Tota 1
8.60d
18.24d
8.00
7.50
Per house
0.14
0.29
6.67
6.67
12.50
12.50
10.00
7.50
6.67
6.67
8.00
7.00
10.00
z
227.27
per house
8.42
Time required to physically operate the tiller.
k Amount of time the tractor motor operated according to the
tractor odometer.
C Amount of fuel used while tractor was in operation (includes
idling time).
d Represents the tractor operating time and fuel utilized to
till one 91-4-m house 63 times, or the tilling site farm (four
91.4-m houses) 15.75 times.

246
Operational costs, i.e. parts and service, over the 2-year testing
period were less than $50. One part had to be rewelded, the oil and
filter were changed, and antifreeze was added to the radiator. Chicken
manure caused rapid corrosion of metal parts, and at^the end of a 2-year
period, the shields covering the tiller's tines were practically rusted
away and replacements were needed. These shields were very important
since they prevented manure from being thrown up into the cages during
tilling and staining the eggs.
Tilling can be incorporated in an Integrated Control Program
insect growth regulators such as CGA 72662 can be applied topically
to manure as a spot treatment in hot spots, areas with thriving popu¬
lations of fly larvae. When tilled into manure, CGA 72662 will prevent
fly resurgence for up to 5 weeks. Similar tests were not attempted
with methoprene because it decomposes so rapidly.
Wood chips can be added to hot spots or wet spots and tilled in to
give manure consistency and aid in the drying process. When chips are
added to semi liquid manure, flat mounds will begin to form if manure is
tilled daily for 1 week. Once chips are added, manure can become
attractive to house flies if moisture levels are right (Kilpatrick and
Schoof, 1959). Tilling should continue in an attempt to reduce moisture
levels below those preferred for fly breeding.
Addition of sand to semi liquid manure causes manure to thicken and
become heavy. The extra weight causes manure to be difficult to handle
and can cause damage to manure spreading equipment. Tilling semi liquid
manure without the addition of a stabilizer is of no value and drying
does not begin until some texture can be given to the manure.

247
Tphyra aenescens Larvae as Predators of Musca domestica Larvae
In the competition study, M. domestica viability was suppressed
significantly by 0. aenescens in all cases (Table 12). This was indica¬
tive of predation but not proof. When larvae of 0. aenescens were reared
in vermiculite, the daily addition of house fly larvae as a possible food
source significantly reduced the mortality rate of 0. aenescens in all
cases (Table 14). Attempts to rear 0. aenescens larvae in vermiculite
failed when no house fly larvae were added. This confirmed that larvae
of 0. aenescens are not cannibalistic, but they are predators of house
fly larvae. Predation by 0. aenescens was also observed during the
study.
Peck (1969) proved that 0. leucostoma, a sister species also found
in poultry manure, is predaceous upon house fly larvae using a vermicu¬
lite study similar to the one I used. Studies indicated that each
larva of 0. aenescens could destroy more than 20 house fly larvae
during the 5~day larval feeding period. Larvae of 0. leucostoma were
found to destroy from 20 (Anderson and Poorbaugh, 1964b) to 30.6 (Peck,
1969) house fly larvae during their developmental period.
Anderson and Poorbaugh (1964b) stated that when house fly larvae
were added to cups of vermiculite containing larvae of 0. leucostoma_,
they were chased out of the vermiculite by the 0. leucostoma larvae.
Ophyra leucostoma larvae killed more house fly larvae than they could
eat and dead larvae attracted more house fly larvae. This enabled
0. leucostoma to increase the number of house fly kills (Anderson and
Poorbaugh, 1964b).

248
Larvae of 0. aenescens in vermiculite became very active when larvae
of M. domestica were added, but did not chase M. domestica larvae as
described by Anderson and Poorbaugh (1964b). The nature of house fly
larvae to congregate around and feed upon the dead of their own kind was
also observed in the 0. aenescens experiments.
When larvae of 0. leucostoma were reared with house fly larvae in
nutrient medium, 0. leucostoma killed as many house flies as when the two
species were reared together in vermiculite. This indicated the prefer¬
ence of 0. leucostoma for house flies over the constituents of the
nutrient medium (Anderson and Poorbaugh, 1964b). This was not the case
with 0. aenescens, as indicated by the competition study.
Morphologica1 Proof that Ophyra aenescens is Predaceous
Seguy (1923) stated that larvae of Ophyra are predaceous but gave
no reasons to substantiate the claim. Morphological evidence was pre¬
sented by Keilin and Tate (1930) who showed that larvae of 0. leucostoma
are predaceous and saprophagous.
The morphological features of the cephaloskeleton of 0. aenescens
are comparable to those of 0. leucostoma. The accessory oral sclerites
and the longitudinal ridges on the venter of the basal sclerite indicate
that larvae of 0. aenescens are predaceous and saprophagous. The cephalo-
skeleton of M. domestica lacks the accessory oral sclerites, but ridges
are present on the venter of the basal sclerite. Larvae with these
characteristics are saprophagous but not predaceous (Keilin and Tate,
1930) .

249
The Value of 0. aenesoens as a Biocontrol Agent
In manure having the desired moisture range, 0. aenesoens may aid
in house fly control if laboratory results are indicative of field re¬
sults. At the tilling site, larvae of Ophyra sp. were only found singly
or in pairs, and never in pockets or hot spots like M. domestica. This
type of behavior would limit the value of Ophyra as a house fly predator.
if a manure management program is drying manure enough to provide
good house fly control, this would also preclude the development of
other díptera such, as Ophyra, whose moisture requirements are similar to
those of M. domestica.
Adults of Ophyra were never noticed at the tilling site except in
light trap catches (Table 40). In large numbers, these flies can be
especially bothersome. Adults are particularly bold and are not intimi¬
dated by the usual methods employed to chase flies from clothing or body
surfaces (P. G. Koehi.er, personal communication).
Rearing Ophyra aenesoens in the Laboratory
Ophyra aenesoens was colonized and reared with the same relative
ease as M. domestica. Life cycle from larva to adult averaged 14 days
at 29.4 C and the average life span for males and females was 16.5 to
20 days respectively. These results were almost numerically the same
as those of Johnson and Venard (1957)-
When kept in the growth chamber at 29.4 C, larvae of 0. aenesoens
raised the temperature of their growth medium to 42.4 C on day 2. The
temperature dropped daily and reached 30.0 C on day 7. When Johnson and
Venard (¡957) found larvae of 0. aenesoens in the field, temperatures of
larvae media ranged from 35.6 C to 41.1 C.

250
High larval densities should be maintained for maximum development
of larvae. Less than 25 larvae per 300 ml of diet resulted in increased
larval mortality. Fortification of the larval diet significantly in¬
creased larval viability.
The Influence of Larvae of Hermetia illucens
on Other Species of Fly Larvae
Larval development of M. domestica_, 0. aenescens, and S. robusta
was not completely suppressed when each species was reared individually
in containers with larvae of H. illucens. Earlier attempts to demon¬
strate complete suppression of other fly species by H. illucens in the
laboratory v/ere inconclusive (Fletcher et al., 1956; Furman et al., 1959).
Soldier fly larvae tend to remain on the bottoms of the rearing
containers and the C.S.M.A. in these areas becomes very slimy. The
slime does not repel the larvae of M. domestica, 0. aenescens, or
S. robusta or otherwise keep them from entering these areas, but it may
somehow inhibit their development.
The order in which the fly species were affected by H. illucens was
M. domestica > S. robusta > 0. aenescens. This was expected for M.
domestica since other experiments indicated that H. illucens would not
allow M. domestica to develop at all (Furman et al., 1959). These re¬
sults were not expected for S. robusta with a larval development period
lasting only 3 days. S. robusta larvae prefer wetter habitats than
M. domestica and perhaps this brought them into closer contact with
H. illucens. Larvae of 0. aenescens were affected least of all by
H. illucens larvae. Mortality rates for H. illucens larvae indicate
that they were not being preyed upon by 0. aenescens. Ophyra larvae
may not be able to penetrate the soldier fly cuticle.

251
In the field, large populations of H. -illucens may totally prevent
other species of flies from breeding in their midst. Control of house
flies on poultry farms with naturally occurring populations of H.
illucens was noted by Tingle et al. (1975) and advocated by Vazquez-
Gonzalez et al. (1962). At the tilling site, populations of house flies
and soldier flies were found living at different depts in the same manure
pack. The depths were determined by moisture level. This is an example
of species packing in which two or more species utilize the same re¬
source without interference (Price, 1975). This unusual situation was
created when wood chips were added to wet manure.
Soldier fly larvae are not popular with poultrymen, especially
after they liquidate manure and then cover the walks when migrating to
dry areas for pupation. This fly is not recommended as a biocontrol
agent in this country. Good manure management techniques should pre¬
vent serious outbreaks of H. -illucens.
The Efficacy of Dimilin as a Feed Additive
When fed to hens at iO ppm, dimilin produced fly mortalities of
Ik.11% in the laboratory and 53.33% in the field. When fed at 1 ppm,
laboratory mortalities were 17.50%. Miller et al. (1975) fed dimilin
to hens at 6.2 to 12.5 ppm and achieved 100% control of flies.
The addition of dimilin to feed at 1 and 10 ppm caused hens to eat
significantly greater amounts of feed and produce eggs at a level that
was numerically but not significantly higher than control hens. Eggs
produced by hens consuming dimilin in the diet have been found to con¬
tain residues (Miller et al., 1975). This, along with the fluorine and
chlorine atoms attached to the dimilin molecule (Kenaga and End, 197^0,
has helped to prevent dimilin from becoming labeled for commercial use.

252
The Efficacy of Methoprene as a Feed Additive
When fed to hens at 10 ppm, methoprene produced fly mortalities of
85.00% in the laboratory and 55.10% in the field. When fed at 1 ppm,
methoprene produced a laboratory mortality of 17-50%. Morgan et al.
(1975) produced 70.9% fly mortality with 5 ppm methoprene diets, but a
mortality of 99-3% required 100 ppm methoprene diets. Low mortalities
produced under field conditions were attributed to the rapid decom¬
position of methoprene (Schaefer and Dupras, 1973). This rapid de¬
composition eliminates methoprene for use as an oral larvicide. Larval
moits are not affected by low concentrations of methoprene and the
critical pupal molt occurs after the larvae have left the area of
methoprene concentration.
Consumption of methoprene caused no significant differences in feed
consumption or egg production except in one case where hens were allowed
to run out of feed. Morgan et al. (1975) found no differences in hen
weights due to methoprene diets.
Methoprene as a Topical Larvicide
2
Methoprene applied topically to manure at 1078 mg/m produced mor¬
talities of 90.0 and 99-0% in laboratory-strain flies 3 days pcst-
2
treatment. Applied at 538 mg/m , 3“day post-treatment mortalities of
73.0 and 96.0% were produced, but this result may not be indicative of
what is occurring in the field.
With a residual of 3 days or longer, methoprene applied topically
would be ideal for spot treatment of house flies in poultry manure.
The granules are in a form that is convenient to use and could be dis¬
pensed from the rear of a rototiller. Tilling would increase the
chances of larvae coming into contact with methoprene.

253
Parasitism rates of pupal parasites have been shown to double in
manure treated with methoprene (Butler and Greer, 197*0, due to the
action of methoprene that prevents pupae from emerging. Used for spot
treating, methoprene could be applied at rates high enough to affect
larval molts as well as the pupal molt. Other investigators found
methoprene to be ineffective as a topical larvicide in poultry manure
(Morgan et a 1., 1975).
Laboratory Studies with CGA 72662
At high concentrations, CGA 72662 kills the early instars of fly
larvae. Only H. illucens and F. can-icularis formed pupae at the initial
levels of CGA 72662 tested, and none of these pupae emerged (Table 27).
An LC^q of 0.A5 ppm was found for house fly larvae. Although
this is well below 1 ppm, the slope of the dosage-mortality curve indi¬
cates that the susceptibility of the house fly population tested to
CGA 72662 is relatively heterogeneous (Hoskins and Gordon, 1956).
CGA 72662 concentrations of 0.25 ppm produced larviform pupae that
eclosed like normal pupae. As concentrations continued to increase,
numbers of larviform pupae increased, and the number of uneclosed
larviform pupae increased (Table 28). At 1.0 ppm five uneclosed larvi¬
form pupae formed in the four treatment replications and no other larvae
or pupae were found. This indicates that at concentrations of 1.0 ppm
and above, larvae were being killed prior to reaching the pupal stage.
CGA 72662, Dimethoate, Dichlorvos, and Ravap
as Topically Applied Larvicides
Effective larva] control lasted for 7 days with Ravap and di¬
methoate, and from 7 to 11 days with dichlorvos. Seven days of control

254
is ail a poultryman can expect to get from compounds that are presently
available (J. F. Butler, personal communication).
The lowest level of CGA 72662 (0.05%) began to break down in 14
days, but the other levels of CGA 72662 were still active 35 days post¬
treatment. CGA 72662 (0.1%) remained active for 35 days even when
tilled into wet manure. Potential resistance problems would result if
CGA 72662 concentrations were not periodically increased to ensure the
production of 100% larval mortalities.
In a liquid form, CGA 72662 may not be as convenient to use as
methoprene granules. However, when used for spot treatment of house
flies in manure, the residual of CGA 72662 would prevent fly activity
for long periods and allow treated areas to dry out. Retreatment would
not be as frequent as with methoprene. CGA 72662, like dimil in, kills
fly larvae in all stages and prevents rapid fly resurgence. Ciba-Geigy
has done residua! testing but the results are unknown at this time.
The Efficacy of CGA 72662 in Water
CGA 726Ó2 added to drinking water of hens at rates of 1.5 to 20.0
ppm produced larval mortality rates in the laboratory house fly strain
of 8O.89 to 100.00% respectively, and was still effective 3 days post¬
treatment. Ciba-Geigy representatives claim that fly control ceases the
day that use of treated water ceases (D. B. Hays, personal communication).
Although the use of a larvicide in drinking water may be considered
impractical by members of the poultry industry (R. H. Harms, personal
communication), Ciba-Geigy feels that all routes of application should
be open to the poultryman (D. B. Hays, personal comnvmication) .

255
Light Traps for Surveying Adult Fly Populations
Each of two light traps at the tilling site caught different numbers
of the monitored fly species from month to month and presented two dif¬
ferent pictures of monthly changes in fly population.numbers. Because
of such variation in catches, light traps are recommended for survey
work but are not considered consistent enough to accurately estimate fly
populations (Pickens et al., 1972).
Driggers (1971) and Pickens et al. (1975) increased fly catches by
lowering traps from ceiling level to ground level. At the tilling site,
traps were hung at ceiling level because ground level traps would have
interfered with the rototiller and would have been covered with manure
as a result of the rototiller.
The yellow trap consistently caught more flies than the black trap.
This was due to the differences in wave lengths emitted by the light
sources of the two traps. Although the traps themselves were two
different colors, it is doubtful that trap color influenced the fly
catch (Mitchell et al., 1975).
Although their influence on fly populations may be questionable,
light traps are looked upon favorably by some poultrymen. Light traps
provide results that the poultryman can see and hear all day long.
Granular Baits for House Fly Control
A combination of Kill'em Fly Killer II (Bomyl) plus Lure'em II
attractant produced the best results in the field although laboratory
testing indicated Kill'em Fly Killer II had a relatively long knockdown
period of ca. 4 hr. Kiil'em Fly Killer (Vapona + ronnel) had the
fastest knockdown, but the lowest fly catches of ali baits tested with

256
the exception of Atroban. Atroban was unattractive to flies in the
field even though laboratory tests indicated it had fly-killing ability
comparable to that of Golden Malrin with Muscamone.
Atroban and Golden Malrin with Muscamone were producing mortality
rates of 87-5 and 100.0% respectively after 40 days of laboratory resi¬
dual testing. Baits field-tested by Bailey et al. (1970) provided 75%
control for half as long. Baits for use on poultry farms do not neces¬
sarily need long residuals. Poultrymen routinely sprinkle baits along
walkways in poultry houses. In the late afternoon while feed is being
distributed to the chickens, walks are swept by a mechanically operated
broom mounted beneath the feed cart.
Wilson and Mulla (1975) observed that bait stations near the peri¬
meters of poultry houses caught flies in nearly a one-to-one ratio and
that bait stations dominated by one sex had catches significantly lower
than those stations conducive to both sexes. Results from field testing
conducted at the tilling site were not always in agreement with the
above observations. The bait catching the most flies attracted signi¬
ficantly greater numbers of females than males.
The addition of Lure'em II attractant to baits resulted in fly
catches significantly higher than those produced by Golden Malrin with
Muscamone. This was not unexpected since other attractants have been
formulated which produce larger catches of flies than Muscamone (Mulla
et al ., 1977).
At the tilling site, flies killed by Muscamone baits were in a 2:1
female to male ratio instead of the 1:1 ratio reported by Carlson and
Beroza (1973). When Lure'em II attractant was added to baits, flies
killed were also in a 2:1 female to male ratio.

257
Granular baits could be used in conjunction with a tilling program
to reduce the numbers in existing fly populations and prevent an increase
in fly numbers due to migrating adults. Attractiveness of baits could
conceivably increase as manure is dried by tilling and becomes less
attractive to flies.
Sugar baits are routinely used by poultrymen to kill adult house
flies. They are easy to use and results are visible in a short period
of time.
Efficacy of Synthetic Pyrethroids as Contact Residuals
Although the house fly LD50 for permethrin (JFU 5819) is above the
acceptable level of 1 ppm, the slope of the dosage-mortality curve indi¬
cates that resistance may be building within the population (Hoskins and
Gordon, 1956). Such resistance has already been reported in mosquitoes
(Priester and Georghiou, 1978). The regression equation presented in
the results section was used to arrive at a tentative LD90 of 45 ppm.
This is below the concentration rates of 500 ppm and 50 ppm tested on
wood, tin, and cement block templates in the laboratory.
Permethrin (JFU 5021A) decomposed in less than 24 and 72 hr on
cement and tin surfaces respectively, but persisted up to 23 months on
wood. This was much longer than could be expected for organophosphorous
compounds (Hansens and Bartley, 1953).
The wooden rafters in poultry houses were logical field application
sites because they are favorite overnight resting places for house flies
(Anderson and Poorbaugh, 1964a). Since synthetic pyrethroids were not
labeled for use in or around poultry houses at the time of the experi¬
ment, selected compounds were sprayed on plywood panels which were hung
in poultry houses at the tilling site after the panels had dried.

258
After flies had been exposed to the panels for ca. 15 min, they
exhibited signs of incoordination and other characteristic symptomology
described by Wouters and van den Bercken (1978). Had they not been caged,
the spasmodic wing fluttering prior to death could have carried flies
far from the panels' surfaces and thus given the impression that the
panels were not efficacious. This may have been what was actually
occurring in the field since dead flies were never seen near the panels.
The synthetic pyrethroid compounds probably persisted at toxic
levels for more than 121 days in the field. Breakdown was attributed
to the disinfectants used to clean the poultry houses during between-
flock sanitation procedures. Once approved for use, synthetic pyrethroids
applied to rafters prior to the arrival of a new flock of chickens
should provide adequate fly control for as long as the flock is housed.
Long term contact residuals like the synthetic pyrethroids would be
a good addition to any fly control program. If applied between flocks
as suggested above, hens, eggs, feed, and water would not be contami¬
nated. The pyrethroids could be applied very thoroughly since the
poultryman would not be concerned with other activities such as daily
flock management. Fly resistance would be of foremost concern. Snono
et al. (1978) stated that ease of permethrin detoxification by insects
may limit its use.
Susceptibility of Endemic Florida Strains of Norther Fowl Mites
to Carbaryl, Malathion, Ravap, and Synthetic Pyrethroids
Laboratory testing showed that the LCr^'s for SO 43775, SBP-1382,
BW 21Z, and Ectiban were higher than those of carbaryl and malathion.
Hall et al. (1978) had similar results except their LC 1s for carbaryl
50
and permethrin, and malathion were greater than mine by approximate

259
factors of 10 and 86 respectively. The mite strains tested by Hall
et al. (1978) were more resistant than the Florida strains, especially
to malathion.
Slopes for all compounds tested were less than ®ne (Table 55).
Carbaryl and malathion had the flattest slopes indicating that concen¬
trations necessary to achieve 100% kill in the field would have to be
much higher than concentrations of the permethrins required to elicit the
same rate of response. The slopes of compounds tested by Hall et al.
(1978) were slightly steeper indicating that the susceptibilities of
their mite strains to the acaricides were more homogeneous than the
Florida strains (Hoskins and Gordon, 1956). In summary, Florida strains
of mites are not as resistant as the strains tested by Hall et al.
(1978), but the shallow slopes of the regression equations indicate
that resistance may be building within the Florida strains.
In field tests, Ravap gave practically 1001 control of mites in
2 weeks. Carbaryl and malathion produced the same results in 4 and 6
weeks respectively. Control with Ravap was probably due in part to
the vaporizing action of Vapona. Loomis et al. (1970) got poor mite
control on hens with a 0.5% carbaryl solution, but Hall et al. (1978)
got 100% control within 24 hr using the same concentration. Several
reports of suspected malathion resistance (Foulk and Matthysse, 1963;
Rodriguez and Riehl, 1963) preceded the confirmation of resistance by
Hall et al. (1978). Many resistance claims are a result of poor appli¬
cation methods (Eleazer, 1978) for which the poultry industry is
notorious. This was not the case with the experiments at the tilling
site, since hens were sprayed more thoroughly than could be expected
under normal conditions.

260
Mite control of 100% was achieved for 36 days with SD 43775 and
BW 21Z on floor-housed hens. Hall et al. (1978) had similar results
for 57 days with much lower concentrations of SD 43775. The reason
for the extended mite control by synthetic pyrehtrin$ may be due to
their ability to persist on the chickens' feathers (Hall et al., 1978).
Since the floor litter was not sprayed, the results of the synthetic
pyrethroid test were particularly interesting. Mites can exist off the
host for 2 to 4 weeks (Cameron, 1938; Baker et al., 1956; Kirkwood, 1963;
Loomis, 1978), and the litter at Chipley, which was compacted and
moderately damp beneath the surface, provided conditions necessary for
mite survival. After short-term acaricides have broken down, mites
hiding in the litter are available to reinfest the birds, but synthetic
pyrethroids persist long enough to break this cycle.
The Effects of Northern Fowl Mites on Egg Production
When 12 strains of laying hens were treated for control of northern
fowl mites, significant differences in production could not be detected
when combined production means were compared by treatment. Although
investigators have claimed that decreases in egg production were due to
northern fowl mites (Cameron, 1938; Metcalf et al., 1962), recent studies
have not been able to corroborate these claims (Loomis et al., 1970;
Eleazer, 1978; DeVaney, 1979). One author suggested that mites be con¬
trolled solely to prevent worker discomfort (Bramhall, 1972).
Comparison of production means of each strain individually revealed
a significant increase in production in one strain due to mite removal.
Five other strains had nonsignificant improvements in production which
compared favorably to the results of Combs et al. (1976).

261
Egg production levels could not be correlated with mite infestation
levels (Figure 45). Perhaps hen performance during the experiment was
not related to mite infestation levels that existed at the time of the
experiment, but to mite infestation levels that existed at some time
prior to the experimental period. Hens infested with mites by DeVaney
(1978) at different points in the laying cycle produced eggs at signifi¬
cantly lower levels 6 to 8 weeks following the onset of infestation.
Hen performance during the experiment at Chipley may also have been due
to the effect of mite infestation levels over time.
During the experiment, mite populations on untreated hens fluc¬
tuated as they must have done prior to and following the end of the
experiment. Therefore, inferences made about egg production and mite
infestation rates may not be entirely correct since the entire picture
is not known. Studies providing for long-term monitoring of egg pro¬
duction and mite infestation rates should provide the data necessary to
determine whether or not a correlation does exist between the two.
Whether it is statistically significant or not, a 1% decrease in
egg production can be costly to a poultry farm owner (R. H. Harms,
personal oommunication) . If a hen lays an average of 240 eggs per year
with a current wholesale value of 5 cents per dozen, a 1% drop in pro¬
duction for just 1 day can cost the owner of a 35,000-bird flock $350.
The first week after birds were treated in Chipley, the production level
of the treated birds was 1% lower than the control birds. One week
later, however, treated birds had increased production by 1.5% and were
again laying at a numerically higher rate than the controls. After the
second treatment, production in the treated birds dropped by 0.64%.

262
This time the production rate was not recovered and after 1 week,
treated birds were laying at a rate which was less than 1% higher than
the control birds. If spraying for mites results in a ]% drop in egg
production for at least 1 week, but cannot guarantee.a \% increase over
and above the original production level, northern fowl mites should be
sprayed, as suggested by Bramhall (1972), only to prevent discomfort to
the labor force.
Long-term compounds like the synthetic pyrethroids could help re¬
duce the number of times per year that hens must be subjected to the
pesticide application stress that occurs when miticides are applied.
A well timed application of a long-term miticide when available, could
break northern fowl mite reproductive cycles and prevent unnecessary
retreatment of hens.
Evaluation of the Hite Rating Systems
The laboratory estimate of northern fowl mite populations was an
attempt to correlate field-estimated mite populations with the actual
numbers of mites on the hens. However, the data obtained by washing
birds are not considered accurate because of the small sample sizes and
the high variability within the samples. Since birds were only washed
once to remove the mite populations, it is not known if all mites were
removed. Also, just one aliquot from each sample was counted.
Since the laboratory estimate was designed to be used only as a
guide, sampling methods that would give statistical confidence were not
employed. The devised conversion factor of 8.13 was also used only as
a guide. Statistically valid conversion factors would require enough
sample numbers to plot a regression line correlating field estimates
and mite numbers for actual mite populations.

263
Mo attempt to correlate field estimates of mite populations with
actual mite numbers present on birds by use of laboratory counting
procedures was found in the literature. The results of all northern
fowl mite field research performed in the past are based on some type
of mite rating system. Hence, the field estimates of northern fowl
mite populations on hens (p. 70) were considered to be the better
estimate of mite numbers and were the values used for statistical
analyses of mite data presented in this dissertation.

CONCLUSIONS
Major conclusions formulated as a result of this research are as
fol1ows:
1. Rototilling was found to be a satisfactory method for drying
wet manure although more time and effort are involved than when a tiller
is used in a routine manure management program.
2. When a tiller is properly incorporated into a routine manure
management program, benefits include fewer manure clean-outs per year,
the increased value of tilled manure as a fertilizer, and reduced costs
for fly control when compared to the use of pesticides.
3. A tiller is not necessary in order to have dry manure on a
poultry farm. Likewise, owning a tiller will not compensate for over¬
all poor management practices.
4. Liquid manure cannot be tilled to a drier state unless a
stabilizer is added. Wood chips aid in the drying process by creating
increased surface area and adding consistency. Builders1 sand makes
manure heavy and difficult to work with.
5. Larvae of Ophyra aenescens are predaceous on house fly larvae
and can kill more than 20 first-instar larvae per day. Large popula¬
tions of Ophyra adults are pestiferous and would be an unwanted
nuisance on a poultry farm.
6. Larvae of Henrietta illueens and Musca domestica can coexist in
larval media in the laboratory and in the field by developing at dif¬
ferent depths i n the media. Drying manure would eliminate both of these
flies.
264

265
7. Methoprene and dimil in added to poultry feed are not effective
as oral larvicides. CGA 72662 in drinking water of hens is an effective
oral larvicide with activity continuing 3 days after cessation of
treatmen t.
8. Methoprene is effective for up to 3 days as a topically
applied larvicide, and would be ideal for use in wet manure since it
is applied in a granular form.
9. CGA 72662 is effective up to 35 days when applied as a topical
larvicide. Fly resurgence can occur within 2 weeks with the use of
commercially labeled organophosphorus larvicides. CGA 72662 persists
long enough to give manure a chance to dry out with no immediate chance
of additional manure 1 iquificat ion due to fly activity.
10. Granular baits may be effective for reducing adult house fly
populations, but probably not as effective as contact residuals. Baits
kill only the flies attracted to them. A contact residual applied to
the rafters of a poultry house would kill all flies coming in to rest
for the night.
11. Northern fowl mites can be controlled with currently labeled
compounds available to poultrymen. Resistance is developing to some
compounds in some areas of the country, but many resistance problems
are due to faulty application methods. Much research is needed in the
area of acaricide application to poultry.
12. Synthetic pyrethroid compounds are effective acaricides with
long residuals. Compounds such as these are necessary to break the
mite life cycle especially when dealing with floor-housed chickens.

266
13. Although northern fowl mites do not significantly affect egg
production, nonsignificant decreases in production due to mites may
result in significant reductions in farm profits.
14. Application of acaricides can cause temporary nonsignificant
drops in egg production that result in monetary losses. Hens in the
latter part of the laying cycle may not be able to regain a rate-of-lay
high enough to recover these losses. More research is needed in this
area .
15. Mite populations do not affect the immediate production level
of hens. However, the affects of present mite populations may be re¬
flected by nonsignificant changes in egg production 6 to 8 weeks in the
future.

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Appendix 1A. Raw data from first Ophyra aenescens adult longevity study.
Day
of Age
Ma1e Morta1ity
No. E %
Fema1e
No.
: Morta1ity
E %
Iota 1
No.
Morta1ity
E *
2
67
-
87
38
-
48.7
*
105
-
67-7
3
10
77
100
17
55
70.5
27
132
85.2
A
-
-
-
6
61
78.2
6
CO
ro
89.O
19
-
-
-
1
62
79.5
1
139
89.7
20
-
-
-
1
63
80.8
1
140
90.3
29
-
-
-
4
67
85.9
4
144
92.9
30
-
-
-
7
74
94.9
7
/
151
97.4
31
-
-
-
A
CO
100.0
4
155
100.0
252

293
Appendix IB. Raw data from second Ophyra aenescens adult longevity study.
Day
of Age
Ha 1 e
No.
Mortality
V O'
¿j 'Q
Fema1e Morta1ity
No. E
Tota 1
No.
Morta1ity
E %
6
-
-
-
1
1
9
1
1
3
10
2
2
1 0
-
-
-
2
3
10
11
1
3
15
1
2
18
2
5
16
16
3
6
30
-
-
-
3
8
26
17
-
-
-
1
•2
27
1
9
29
18
3
9
45
-
-
-
3
12
39
20
4
13
65
2
5
45
6
18
58
21
1
14
70
-
-
-
1
19
61
22
2
16
80
1
6
55
3
22
71
23
1
17
35
-
-
-
1
23
74
24
-
-
-
1
7
64
1
24
77
25
-
-
-
1
8
73
1
25
81
27
1
18
90
1
9
82
2
27
CO
28
1
19
95
1
10
91
2
29
94
30
1
20
100
-
-
-
1
30
97
31
-
-
-
1
11
100
1
31
100

294
Appendix 1C. Raw data from third Ophyra aenescens adult longevity study.
Ma1e Morta1 ity Fema1e Mortai ity Tota 1 Morta1ity
1 Age
No.
E
%
No.
E
%
No.
E
%
3
_
_
_
1
1
0.5
1
1
0.3
4
1
1
0.6
1
2
0.9
2
3
0.8
6
1
2
1.1
-
-
-
1
4
1 .0
8
1
3
1.7
1
3
1 .4
2
6
1.5
9
2
5
2.8
-
-
-
2
8
2.0
12
1
6
3.3
-
-
-
1
9
2.2
13
6
12
6. 6
-
-
-
6
15
3.8
15
9
21
11.6
3
6
2.8
12
27
6.8
16
28
49
27.2
5
11
5.1
33
60
15.1
17
25
74
41 .1
8
19
8.8
33
93
23.4
18
21
95
52.8
13
32
14.7
34
127
32.0
19
21
116
64.4
8
40
18.4
29
156
39.3
20
4
120
66.7
12
52
24.0
16
172
43-3
21
12
132
72.3
20
72
33.2
32
204
51.4
22
11
143
79.4
36
108
49.8
47
251
63-2
23
8
151
83.9
30
138
63.6
38
289
72.8
24
11
162
90.0
20
158
72.8
31
320
80.6
25
1
163
90.6
21
179
82.5
22
342
86.1
26
4
167
92.8
8
187
86.2
12
354
89.2
27
4
171
95.0
6
193
88.9
10
364
91 .7
28
2
173
96.1
3
196
90.3
5
369
92.9
29
2
175
97.2
1
197
50.8
3
372
93-7
30
1
176
97.8
4
201
92.6
5
377
95.0

255
Appendix 1C. Continued.
Day
of Age
Ma 1 e
Morta1ity
Fema1e Morta1ity
Tota 1
Morta1ity
No.
Z
%
No.
E
%
No.
£
%
31
1
177
98.3
-
-
-
1
CO
PO
95.2
32
-
-
-
4
205
94.5
4
382
96.2
33
1
178
98.9
1
206
94.9
2
V^>
CO
-C-
96.7
34
1
179
99.4
3
209
96.3
4
OJ
CO
oo
97-7
35
1
180
100.0
1
210
96.8
2
390
98.2
36
-
-
-
1
21 1
97.2
1
391
98.5
40
-
-
-
1
212
97.7
1
392
98.7
42
-
-
-
1
213
98.2
1
393
99.0
45
-
-
-
1
214
98.6
1
394
99.2
47
-
-
-
3
217
100.0
3
397
100.0

3
5
6
7
3
9
10
11
12
13
14
15
16
17
13
19
20
21
22
24
25
26
27
236
1D. Raw data from fourth Ophyra aenesoens adult longevity study.
Ma1e Morta1 ity Fema1e Mortality Tota 1 Morta1ity
No.
£
O/
'O
No.
v
Lj
%
No.
£
%
2
2
0.4
-
-
*
2
2
0.2
12
14
3.2
34
34
8.0
46
48
5.6
32
46
10.5
43
77
18.1
75
123
14.2
2
48
11.0
7
84
19.7
9
132
15.3
-
-
-
5
89
21 .0
5
137
15.9
10
58
13.2
19
108
25.4
29
1 66
19.2
5
63
14.4
3
111
26.1
8
174
20.1
7
70
16.0
1
112
26.3
8
182
21 .1
20
90
20.5
3
115
27.0
23
205
23-7
27
117
26.7
7
122
28.6
34
239
27.7
49
1 66
37.9
8
130
30.5
57
296
34.3
87
253
57.8
23
153
35.9
110
406
47.0
72
325
74.2
32
185
43.4
104
510
59.0
27
352
80.4
33
218
51 .1
60
570
66.0
37
389
88.8
39
257
60.3
76
646
74.8
17
406
92.7
24
281
66.0
41
687
79.5
13
419
95.7
37
318
74.6
50
737
85.3
5
424
96.8
15
333
78.2
20
757
87.6
7
431
98.4
33
366
85-9
40
797
92.2
-
-
-
17
383
89-9
17
814
94.2
-
-
-
14
397
93.2
14
828
95.8
-
-
-
7
404
94.8
7
835
96.6
-
-
-
6
410
96.2
6
841
97-3

257
Appendix ID, Continued.
Day
of Age
Ma1e Morta1ity
No. Z X
Femaie Morta1ity
No. £ %
Tota 1 Morta1ity
No. Z %
28
3
43 4
99.1
4
414
97.2
7
/
848
98.1
29
1
435
99.3
2
416
97.7
3
851
98.5
30
-
-
-
2
418
98.1
2
CO
\J"1
CO
98.7
32
2
437
99.7
3
421
98.9
5
858
99-3
36
1
OO
100.0
3
424
99.5
4
862
99.8
39
-
-
-
2
426
100.0
o
L
864
100.0

Appendix 2. Raw data from CGA 72662 laboratory studies.
Fly Species
CGA 72662
Concent rat ions(%)
M. domestica control
0.06
0.05
* 0.04
0.03
7/1 0a
0/10
0/10
0/10
0/10
7/10
0/10
0/10
0/10
0/10
9/10
0/1 0
0/10
0/10
0/1 0
6/10
0/10
0/10
0/10
0/10
Z 29/1+0
0/40
0/40
0/40
0/40
% Morta1ity 27-5
100.0
1 00.0
100.0
100.0
H. illucens control
0. 10
0.075
0.050
0.025
10/10
0/1 0
2/10
0/10
6/10
10/10
0/10
1/10
3/10
5/10
10/10
0/10
0/10
3/10
6/10
10/10
0/10
2/10
0/10
2/10
E 40/40
0/40
5/40
6/40
19/40
% Morta1ity 0.0
100.0
87.5
85.0
52.5
S. robusta control
0.10
0.075
0.050
0.025
8/10
0/10
0/10
0/10
0/10
9/10
0/10
0/10
0/10
0/10
10/10
0/10
0/10
0/10
0/10
10/10
0/10
0/10
0/10
0/10
Z 37/AO
0/40
0/40
0/40
0/40
% Mortality 7-5
100.0
100.0
100.0
100.0
P. regina control
0.10
0.075
0.050
0.025
10/10
0/10
0/10
0/10
0/10
8/10
0/10
0/10
0/10
0/10
10/10
0/10
0/10
0/10
0/10
10/10
0/10
0/10
0/10
0/10
Z 33/40
0/40
0/40
0/40
0/40
% Morta1ity 5-0
100.0
1C0.0
100.0
100.0
298

295
Appendix 2. Continued.
CGA 72662
Fly Species
Concent rat ions{%)
0. aenesasns control
0.10
0.075
- 0.050
0.025
8/10
0/10
0/10
0/10
0/10
8/10
0/10
0/10
0/10
0/10
8/10
0/10
0/10
0/10
0/10
8/10
0/10
0/10
0/10
0/10
Z
32/40
0/40
0/40
0/40
0/40
% Morta1ity
20.0
100.0
100.0
100.0
100.0
F. canioularis control
0.10
0.075
0.050
0.025
9/10
4/1 0
2/1 0~
0/1 o1
5/1 0T
6/10
1/1 o"
3/10
5/1 o"
6/1 0_
5/10
1 /1 o'c
7/io*
2/1 O7
5/1 o"
10/10
3/10*
5/10“
3/10T
3/1 0~
Z
30/40
9/40
17/40
10/40
19/40
% Mortaiicy
25.0
77.5
57.5
75.0
52.5
3 Represents the
number
of
pupae that
ec1osed
unless
otherwise
stated.
k Values are the
number
of
pupae that
formed.
Pupa 1
morta1ity
was 100% in all cases
except
the
controls.
Indicates the
presence
; of
dead larvae.
Indicates the presence of larviform pupae.

Appendix 3. House flies killed in Farnam bait field trial.
House
No.
1
Kill1 em Fly Killer 1
I Ki111em
2
Fly Bait
3
Kill'em Fly Killer
o
o
o
o
o
o
1
6
11
33
38
1
0
3
8
8
7
10
1
2
1
6
8
0
1
0
0
3
7
9
2
0
0
0
Total
by
Sex 27
37
42
49
2
2
Grand
Tota 1
64
91
4
1+5
2+5
3+5
Ki 111em Fly Killer i
1 K i 1 1 'em
Fly Bait
Kill'em Fly Killer
House
+
+
+
Mo.
Lure
'em II
Lure
'em II
Lure
'em II
o
o
o
o
c
o
2
92
280
172
288
17
52
4
72
144
63
157
12
62
2
97
285
43
81
0
10
4
81
215
21
104
1
3
Tota 1
by
Sex 342
924
299
630
30
127
Grand
Total 1266
929
157
300

301
Appendix 3- Continued.
4
6
7
House
SX-70
Fly Ba i t
Golden Malrin^.^
w/o Muscamone
Golden Ma 1 r i n
with Muscamone
No.
o
o
o
o
* o
o
1
53
45
31
50
5
14
3
39
63
17
20
6
21
1
2
1
3
2
1
0
3
3
18
1
2
0
2
Tota 1
by
Sex
102
127
52
74
12
37
Grand
Tota 1
229
126
49
4+5
6+5
7
House
No.
SX-70 Fly Bait
+
Lure 'em II
Golden Malrin_^
w/o Muscamone
+
Lure 1em II
Golden Ma1rin
with Muscamone
o
o
o
o
o
o
2
12
26
17
49
15
18
4
17
44
17
61
8
16
2
5
20
3
8
2
1
4
5
32
0
5
1
2
Tota 1
by
Sex
39
122
37
123
26
37
Grand
Tota 1
1 61
160
63

Appendix 4. The probabi 1 iti es, probits, log doses, and lower and upper
fiducial limits from the probit analysis of SBP-1382
dosage-mortality data.
0(3 S
PROS
OR03IT
LOOSE
LCWE R
UPPER
1
0.0 1
2.67
-11.02
-17.62
— 9.o5
2
0.02
2 • 95
- 1 0.67
-16.51
-9.44
~7
0.03
3.12
— 10.46
- 1 5*. 96
-9.3 1
4
0.04
3.25
-10.29
-15.47
-9.2 1
5
0.05
3.36
-10.16
-15.05
— 9.12
5
3.06
3.45
-10.05
-14.75
— 9.0 o
7
C . 07
3.52
-9.95
-14.45
-3.99
a
0.03
3.59
-9.36
-14.19
-3.93
9
0.09
3 . o ó
-9.76
-13.95
- 3.83
l 9
0.10
3.72
-9.71
-13 .73
-3.34
l 1
0.15
3.96
-9.40
- 12.32
-3.64
1 2
0.2 0
4.16
— 9.15
-12.11
-3.43
13
0.25
4.33
-3.94
-11.50
-3.33
1 4
0.30
4.46
-3.76
- 10.95
-3.20
1 5
0.3 o
4.6 1
— 3.53
-10.46
-3.07
1 5
0.40
4.75
— 3.42
-9.9 9
-7.94
1 7
0.45
4.37
-3.26
-9.60
-7.79
1 6
0 » o 0
5.00
-3.10
—9.16
-7 ,o3
1 9
0.55
5.13
-7.94
-3.73
-7,44
20
0.60
5.25
-7.73
— 3.44
-7.20
2 t
0.65
5.39
-7.62
— 3.16
— 6.90
22
0.7 0
5.52
-7.44
-7.92
— ó . 51
2 3
0.7 5
5.57
-7.25
-7.71
-6.04
2 4
0.30
6.34
-7.04
-7.52
-5.47
2 5
0.3 5
6.04
— 6.3 0
-7.33
-4.79
£6
0.90
6.28
-6.49
-7.11
-3.90
27
0.91
6.34
-6.42
-7.06
— 3.69
2 8
0.92
6.4 1
-5.34
-7.00
— 3,40
29
0.93
6.46
-5.25
—6 .94
-3.19
3 O
0.94
6 . o 5
-6.15
-6.63
-2.90
3 1
0.95
6 . ó 4
— 6.04
-6.30
-2.57
3 2
0.9 6
6.75
-5.90
-ó . 7 2
-2.13
33
0.97
6.33
-5.74
- 6 . ó 1
-1.70
_/ -r
0.9 3
7.05
-5.52
— ó » 4 3
— 1 ,0o
3 5
0.99
7.33
-5.13
-6.26
-0.04
302

Appendix 5. The probabilities, probits, log doses, and lower and upper
fiducial limits from the probit analysis of BW 21Z dosage-
morta1ity data.
G d S
?RC3
PR0 3 I T
LOOSE
LOWER
U ?°£R
1
0.01
2.67
-10.71
- 1 1.37
-10.13
"5
0.02
2.95
-10.43
-11.1-4
-9.9 1
3
0.03
3.12
-10.25
-11117
-9.7 -3
4
0.04
3.25
-10.11
-10.97
-9.67
5
0.05
3.36
-10.00
-1C .31
— 9.5 4
6
0.06
3.43
-9.9 1
-10.67
-9.52
7
0.07
3.52
-9.33
— 10.55
-9.46
3
0.03
3.59
-9.75
-10.44
-9.40
9
0.09
3.66
-9.69
-10.34
-9.35
10
0.10
3 .72
-9.63
-10.25
— 9.30
1 1
0.15
3.96
-9.37
-9.37
-9.10
12
0.20
4.16
-9. 17
-9.53
-3.94
1 3
0.25
4.33
-9.00
-9.33
-3.30
1 4
0.30
4.43
-3.34
-9.12
— 3 » ó 6
1 5
0.3o
4.61
-3.70
-3.93
-3.53
1 5
0.40
4.75
-3.56
— 8.75
-3.39
1 7
0.45
4.37
-3.43
— tt . 6 1
-3.24
1 3
0.50
5.00
-3.3 0
-3.47
-3.03
19
0.55
5.13
-3.17
- a. 3 5
-7.91
20
0 . Ó0
5.25
- 3.04
-8.23
-7.73
2 1
0.65
5.39
-7.90
-3.11
-7.54
2 2
0.70
5.52
-7.75
-8.00
-7.33
23
0.75
5.57
-7.50
-7.37
-7.10
2a.
0.30
5.34
-7.43
-7.7 4
-6.35
25
0.35
6.04
-7.23
-7.53
— 6.55
2 ó
0.90
6.23
-6.97
— 7.33
— 6.17
27
0.91
6.34
-6.91
-7.34
—6.03
23
0.92
6.41
-6.35
-7.2 )
-5.9,3
29
0.93
6 » 43
-6.77
-7.2 3
-5.37
30
0.94
6.55
— ó » 6 9
-7.17
-5.75
3 1
0.95
6.64
— ó . 6 0
-7.10
— 5.0 1
32
0.96
6.75
-6.49
-7.01
— 5.44
3 3
0.97
6.33
-ó.35
— 6.91
-5.24
34-
0.93
7.05
-6.17
-6.73
-4.97
35
0.9 9
7.33
-5.39
— ó . 56
— 4.0 5
303

1
2
3
4
5
6
7
3
9
1 O
1 1
1 2
13
1 4
1 5
1 6
1 7
1 3
I 9
20
21
22
23
24
2 5
28
27
28
29
30
J 1
82
33
34
35
The probabilities, probits, log doses, and lower and upper
fiducial limits from the probit analysis of SD 43775
dosage-mortality data.
PROS
PROSIT
LO 300
LOWER
UPPER
0.01
2.57
-10.07
-11.03
-9.49
0.02
2.95
-9.79
-10.70
-9.23
0.03
3.12
-9.62
-10.46
-9.14
0.04
3.25
-9.49
-10.23
-9.04
0.05
3 • 36
-9.38
-10.13
— 3.95
0.06
3.45
-9.29
-10.01
-3.33
0 .07
3.52
-9.21
-9.90
-8.82
0.03
3.59
-9.14
-9.30
-8.76
0.0 9
3.56
-9.03
-9.71
-3.71
0.10
3.72
-9.02
-9.63
— 8 . 66
0. 15
3.96
-3.77
-9.29
-3.46
0.20
4.16
— 8.53
-9.02
-3.31
0.25
4.33
-3.4 1
-S.30
-3.17
0.30
4.48
-3.26
— 3.60
-3.0 +
0.35
4.61
-3.12
— 3.41
-7.92
0.40
4.75
-7,93
-8,24
-7.30
0.45
4.37
-7.36
- S . 0 3
-7.63
0.50
5.00
-7.73
-7.93
— 7.56
0.55
5.13
-7.50
-7.73
—7.43
0.60
5.25
-7.43
— 7.65
-7.29
0.65
5.39
-7,34
-7.51
-7.13
0.7 0
5.52
-7.20
-7.33
— 6.96
0.75
5.67
-7.05
-7.25
-6 . 77
0.80
5.84
-6.33
-7.10
— ó . 5 5
0.8a
6.04
— 6.69
-6.94
-6. 29
0.90
6.23
— 5.44
-6.73
— 5.95
0.9 1
6.34
-5.33
— ó » 6 9
-5.37
0.92
6.41
-6.32
— 6.63
- 5 • 7 9
0.93
6.43
-5.25
-6.53
-5.69
0.94
6.55
-6.17
— 6.51
— 5.53
0.95
6.64
-6.08
-6.44
— 5.45
0.96
6.75
-5.97
— 6.35
-5.33
0,97
6.33
-5.34
-6.25
— 5.13
0.96
7.05
- 5 . ó 7
-6.11
— 4.89
0,99
7.33
-5. 39
-5.90
-4.51
304

Appendix 7.
The probabi i ities , probits, log doses, and
fiducial limits from the probit analysis of
dosage-mortality data.
lower and upper
ICI Ectiban *^
DBS
PRG3
PRC51T
LOGOOS
LOWER
UPPER
1
0.0 1
2.67
- i 1.73
-12.73
- 1 1 .03
2
0.02
2.95
-11.37
-12.31
-10.79
3
0.03
3.12
-11.14
- 1 2*. 0 0
-10.60
4
3.04
3.25
-10.97
- 1 1 . 7 5
—13.46
5
0.05
3.36
-10.32
-11.59
-10.35
6
0.06
3.45
-10.71
-11.43
-10.25
7
0.07
3.52
- 10.60
-11.30
—10.16
3
0.03
3 .59
-10.51
-11.17
-10.09
9
0.09
3 • ó 6
-10.42
-11.Oó
-10.02
10
0.10
3.72
—10.54
-10.96
-9.95
1 1
0.15
3.96
-10.02
—10.54
-9.68
1 2
0.20
4.15
-9. 76
- 1 0.20
— 9.46
13
0.25
4.33
-9.54
-9.92
-9.27
l 4
0.30
4.43
-9.34
-9.67
-9.10
15
0.35
4.61
-9.15
— 9.45
-3.93
16
0.40
4.75
-3.93
-9.24
-3.76
1 7
0.45
4.37
-3.8 1
-9.04
— 3.60
13
0.50
5.00
— 8 . ó 4
— 3.3 ó
-3.42
19
0.55
5.13
-3.47
— 8.69
-6.24
20
0.60
5.25
-3.31
-S. 52
-3.05
2 1
0.65
5.39
-3.13
-3.35
-7.84
22
0.70
5.52
-7.95
-8.18
— 7 * 6 1
23
0.75
5.67
-7.75
-8.01
-7.37
24
0.30
5.34
-7.52
-7.32
-7.03
25
0.35
6.04
-7.27
-7.53
—6.75
2 5
0.90
6.23
-6.94
-7.33
-5.33
27
0.91
6.34
-6.36
-7.25
-5.23
23
0.92
0.41
-6.73
- 7 . i 9
-5.11
29
0.93
6.43
— 6.63
-7.12
-5.99
30
0*94
6.55
— 6.58
-7.03
— 5 • o o
31
0.95
6 • 64
-5.46
— 6.9 _>
-5.70
32
0 • 96
6.75
-6.32
-6.32
— 5.51
33
0.97
6.38
— 6.15
— 6.68
— 5.23
34
0.93
7.05
-5.92
— 6,4 9
— 4 . 9 9
3d
0.99
7.33
— 5.55
-6.20
— 4.50
305

i I X
33
1
2
3
4
5
6
7
a
9
19
1 1
1 2
13
14
15
1 5
1 7
i 3
1 9
20
21
22
23
¿■i
25
26
2 7
23
29
30
3 i
32
33
3 4
35
The probabilities, probits, log doses, and lower and upper
fiducial limits from the probit analysis of carbaryl
dosage-mortality data.
PR C 3
PftOBIT
LQGDOSE
LOWER
UPPER
9.01
2.67
-13.19
-23.54
-15.73
0.02
2.95
-17.23
-22.01
-15.09
0.03
3.12
-16.64
- 2 1*. 0 4
-14.65
0.04
3.25
-16.19
-20.31
-14.32
0.05
3.36
-15.82
- 19.71
-19.05
0.9 ó
3.45
-15.51
-19.21
-13.32
0.07
3.52
-15.23
— 1 3.77
-13.61
0.03
3.59
-14.96
-13.37
— 1 O . 4 3
0.09
3 .66
—14.76
-18.01
-13.27
0.13
3. 72
- 14.56
—17.68
-13.11
0.15
3.96
-13.70
- 16. i l
- 12.43
9.20
4.16
-13.02
-15.23
-11.97
0.25
4.33
-12.44
-14.31
- 1 1 . 53
0.30
4.43
-11.92
-13.50
-11.12
9.35
4.61
-11.44
-12.75
-10.73
0.40
4.75
-10.98
-12.06
-13.34
0.45
4.37
-10.53
-11.41
-9.95
0.50
5.00
-10.09
-10.32
-9.51
0.55
5.13
— 9 • ó 6
-10.23
-3. 03
0.60
5.25
-9.21
-9.79
-a .43
0.65
5 . o ^
— 3.75
-9.34
— 7 > 85
0.70
5.52
-6.27
-3.91
-7.14
0.7 5
3.67
-7.75
— 3 .43
—6.35
0.30
5.34
-7.16
-3.02
-5.44
0.85
ó .04
-5.49
-7.50
-4.33
0.90
6.23
— 5.63
-6 .36
-3.0 2
0.9 1
6.34
-5.43
-6.7 0
-2.69
0.92
6.41
-5.20
— 6.54
-2.33
0.93
6.43
— 4.96
-6.35
— 1 .94
0.94
5.55
-4.68
-6.15
- 1.49
0.95
6.64
-4 .37
-5.92
- C . 9 9
J.9 6
.6.75
-4.00
-0.65
-0.40
0.9 7
6.38
— 3.55
— 5.31
0.33
0.93
7.05
-2.95
-4 .3 7
1 .33
0.99
7.33
-1.99
— 4 • 1 3
2.33
306

Appendix 9. The probabilities, probits, log doses, and lower and upper
fiducial limits from the probit analysis of malathion
dosage-mortality data.
as
P9 03
PROSir
LCOCSE
LOWER
UPPER
i
0.0 1
2.57
-13.52
-15.35
-12.43
2
0.02
2.95
-12.95
-14.59
-11.97
3
0.03
3.12
-12.59
- 14.12
— 1 1 .63
4
0.04
3.25
-12.32
-13.74
-11.46
5
0.05
3.36
-12.10
—13.45
- 1 l .23
6
0.05
3.45
-11.91
-13.20
-11.13
7
0.07
3.5 2
-11.75
-12.93
-10.99
a
0.03
3.59
-1 I .60
-12.73
-10.83
9
0.09
3.66
- 1 1.47
-12.61
-10.77
1 0
0.10
3.72
-11.34
— 12.44
—10.67
11
0.15
3.96
-10.33
-1 1.77
-10.25
1 2
0.20
4.15
-10.42
-11.23
-9.91
1 13
0.25
4.33
-10.oa
-10.73
-9.61
l 4
0.30
4.48
— 9.7 ó
- 1 0.33
— 9.3b
1 5
0.35
4.61
- 9.47
-10,01
-9.39
1 6
0.4 0
4.75
-9.20
- 9 . ó 7
-3.34
1 7
0.45
4.37
-3.93
-9.3a
-3.59
1 3
0.50
5.00
-0.67
-9.04
— 3.3.3
19
0.55
o . 1 3
-3.41
— 3.75
— 3.06
2 0
0.50
5.25
-3.14
- 8.4 7
-7.77
2 1
0.55
5.39
-7.36
-3.20
— 7.45
22
0.70
5.52
-7.57
— 7.93
-7.11
23
0.75
5.67
-7.26
-7.64
-6. 7 8
? 4
0.3 0
5 • 34
-5 .91
— / .04
-6 . ? 7
2 5
0.6
6.04
— ó » b 0
-6.99
- 5.7 5
2 n
0.90
6.23
-5.00
— 6.57
-5.09
2 7
0.91
6.34
-5.37
-6.4 7
— 4.92
2 a
0.92
6.41
-5.74
— 6,3 ó
— 4 * / 4
2 9
0.93
6.43
— 0.59
-c.23
— 4.54
30
0.94
6.55
-5.42
-6.13
-4,33
3 1
0.95
6.54
-5.24
-5.95
-4.03
32
0.95
6.75
-5.0 1
-5.77
— . ! 3
.33
0 . 9 7
6.33
-4,74
— 5.55
- 3 , a 2
3 4
0.93
7.05
-4.33
-5.25
-2.94
35
0.99
7.33
-3.31
-4.30
-2.13
307

BIOGRAPHICAL SKETCH
Jerome Adkins Hogsette, Jr., was born on MarclT 5, 1945, in Miami,
Florida. He graduated from North Miami Senior High School in June of
1963, and received an A.A. from Miami-Dade Junior College in April of
1966. In September of 1966, he entered the University of Florida,
but joined the U.S.A.F. in May of 1968 prior to the completion of his
bachelor's program. During a 4-year tour of duty in the Air Force,
the author was involved with sanitary inspections of food and food
service facilities, and health care programs for sentry dogs and
animals belonging to military personnel.
After separating from the Air Force in March of 1972, Jerome
re-entered the University of Florida and received a Bachelor of
Science in Poultry Science in December of 1973- In August of 1975,
he received a Master of Science in Agriculture with a major in
poultry science and a minor in entomology. In January of 1976, the
author began working on a Ph.D. program in entomology, which he is
presently completing. Jerome is a member of the Entomological
Society of America.
308

I certify that I have read this study and that in my opinion
it conforms to acceptable standards of scholarly presentation and
is fully adequate, in scope and quality, as a dissertation for
the degree of Doctor of Philosophy.
J err// F. Bu"tp4r\ Cha i rman
Professor of Entomology and Nematology
I certify that I have read this study and that in my opinion
it conforms to acceptable standards of scholarly presentation and
is fully adequate, in scope and quality, as a dissertation for
the degree of Doctor of Philosophy.
P. G. Koehler
Associate Professor of Entomology and Nematology
I certify that I have read this study and that in my opinion
it conforms to acceptable standards of scholarly presentation and
is fully adequate, in scope and quality, as a dissertation for
the degree of Doctor of Philosophy.
iQ lO. \¿*SUL
D. W. Hall
Associate Professor of Entomology and Nematology
I certify that I have read this study and that in my opinion
it conforms to acceptable standards of scholarly presentation and
is fully adequate, in scope and quality, as a dissertation for
the degree of Doctor of Philosophy.
(3. Qk
/
1 ¿<2-
C. R. Douglas t/
Associate Professor of Poultry Science

This dissertation was submitted to the Graduate Faculty of the
College of Agriculture and to the Graduate Council, and was
accepted as partial fulfillment of the requirements for the
degree of Doctor of Philosophy.
December, 1979
Dean, Graduate School

UNIVERSITY OF FLORIDA
3 1262 08554 0788




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