Citation
The black flies (Diptera : Simuliidae) of Florida and their involvement in the transmission of Leucocytozoon smithi to turkeys

Material Information

Title:
The black flies (Diptera : Simuliidae) of Florida and their involvement in the transmission of Leucocytozoon smithi to turkeys
Creator:
Pinkovsky, Dennis Drew, 1947- ( Dissertant )
Butler, J. F. ( Thesis advisor )
Fairchild, A. G. B. ( Reviewer )
Walker, T. J. ( Reviewer )
Dame, D. A. ( Reviewer )
Forrester, D. J. ( Reviewer )
Fry, Jack L. ( Reviewer )
Place of Publication:
Gainesville, Fla.
Publisher:
University of Florida
Publication Date:
Copyright Date:
1976
Language:
English
Physical Description:
xiii, 331 leaves : ill. ; 28 cm.

Subjects

Subjects / Keywords:
Black flies ( jstor )
Cocoons ( jstor )
Creeks ( jstor )
Eggs ( jstor )
Female animals ( jstor )
Larvae ( jstor )
Pupae ( jstor )
Rivers ( jstor )
Species ( jstor )
Streams ( jstor )
Dissertations, Academic -- Entomology and Nematology -- UF
Entomology and Nematology thesis Ph. D
Flies -- Florida ( lcsh )
City of Gainesville ( local )
Genre:
bibliography ( marcgt )
non-fiction ( marcgt )

Notes

Abstract:
Immature and adult black flies (Diptera: Simuliidae) were collected in Florida over a period of three years. Eighteen species of black flies including ten which are new records for the State were found to occur in Florida. Records of black flies from 192 locations in 50 counties are included. Biological information is provided for each species together with data on distribution, seasonal occurrence, stream ecology and species associations. Keys to the Simuliidae of Florida are provided and structures are illustrated. Representative specimens have been deposited in the Florida State Collection of Arthropods and in the United States National Museum, Washington D.C. Transmission investigations have incriminated three species, Simuliwri congareenarum, S. meridionale, and S. slossonae, as vectors of Leucocytozoon smithi to turkeys in Florida. On nineteen occasions L. smithi was transmitted to domestic turkeys by the bites of infected black flies.
Thesis:
Thesis (Ph. D.)--University of Florida, 1976.
Bibliography:
Includes bibliographic references (leaves 296-318).
General Note:
Typescript.
General Note:
Vita.
Statement of Responsibility:
by Dennis Drew Pinkovsky.

Record Information

Source Institution:
University of Florida
Holding Location:
University of Florida
Rights Management:
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:
022389079 ( alephbibnum )
ACZ5866 ( notis )
02526789 ( oclc )

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THIE BAICK FLIES (DIPTERA:: SIHULLIIDAE) OF FLOIDAT, AN1\D 'THEIR
INVOLVEMENT INi THE TRANSMISSION OF'
L~eucocytozoon smith TO TURKEYS








By

DENNIS DREW PINKOVSKY


A D)ISSERTATIOJN PRESENTED TO THE GRA.DUAT~ COUNCIL OF
THE UNIVERSrTY OF FLOREDAl
IN PARTIAL FULFILLMENTT UF THE REQUIREMENTS FOR\ TbE
DEGREE OF DOCTOR OF PHILOSOPHti












UNIZVERSITY OF FLO;:rIDA
















ACKNOWLEDGEMIENTS


I sincerely thank Dr. J.F. Butler, my Committee Chairman, for his

constructive suggestions and support during my research endeavors and

for his valuable, critical review of my dissertation.

To Dr. D.F. Forrester I extend my deep appreciation for the advice

and guidance he enthusiastically offered during my transmission studies

and for the equipment and facilities he generously allowed me to use.

To all the mem~btrs of my Ph.D. Committee I express appreciation

for their critical appraisal of my dissertation and their helpful

comments.

I wish to thank Dr. E.L. Snoddy and Dr. G.E. Shewell for examining

black fly specimens which I sent from Florida and for the determinations

rendered.

I express my gratitude for the opportunity to examine black fly

specimens collected in Florida which were made available to me by

numerous individuals and institutions.

To the staff at the U.S. Department of Agriculture Insects Affecting

Man Laboratory, Gainesville, I extend my appreciation for the use of equip-

ment and for advice during my research and academic studies.

I am grateful to Dr. E.V. Komarek and the staff at the Tall Timbers

Research Station and L..E. pjilliams, D.H. Austin, and T. Peoples of the

Florida Game and! Fresh Water Fish~ Commission for the use of facilities

and other assistance during my collecting trips.









I wish to also thank the State of Florida Division of Recreation

and Parks for collecting permits and the opportunity to gather specimens

at the beautiful State Parks around Florida.

I extend my deep gratitude to the U.S. Air Force Institute of

Technology Civilian Institutions Program for financial support and for

the opportunity to attend the University of Florida and pursue my Ph.D.

degree.

To P. Humphrey, D. Young, L. DuBose and especially T. DiNuzzo I

express my sincere thanks for good-spirited support during my research.


iii




















































































General Comments .


Introduction to the Black Fly Keys. ..... .


A key to the larvae of the black flies of
Florida . . . . . *

A key to the pupae of the black flies of
Florida . . . . . . .


TABLE OF CONTENTS


ii


vi


vii


ziii


ACKNOWLEDGEMENTS. ....


LIST OF TABLES. ...c.


LIST OF FIGURES .....


AB STRACT. ........


CHAPTER


I. INTRODUCTION. ..........


II. LITERATURE REVIEW ........


Simuliidae. ...........


Taxonomy and Distribution. .
Bionomics. .........

Damage .. . .. . .. .
Control. ..........


Florida Ecological Habitats ...


Leucocytozoon smith. ......


III. MATERIALS AND METHODS ,.....


Black Fly Survey. ........


Leucocy-tozoon smithi Transmission


IV. RESULTS AND DISCUSSION. .....


Simuliidae. ...........


. .


. . .


. . .


. . .


. . .
. . .
. . .
. . .


. . .


. . .


. . .


. . .


. . .


. . .


. . .


. . .









TABLE OF CONTENTS
(Continued)


Page


A key to the adult male black flies of
Florida . . . . . . . .
A key to the adult female black flies of
Florida . . . . . . . . .

Introduction to the Individual Species Sections...
Chephia (Cnephia) ornithophilia Davies, Peterson,
and Wood . . . . . . . .
Simutiumr (Byssodon) meridionaie Riley. ......
Simulium (Bt~ssodon) slo~ssonae D~yar and Shannon ..
Sirmuium (Eurshnutium) cong~areenatum(Dyar and
Shannon) . . . . . . . . .
Simutium (Phosterodoros) dixieinse Stone and Snoddy
Simutium (Phosterodoros) haysi Stone and Snoddy..
Simu~ium (Phosterodoros) denningsi Malloch.. ..
Simutlium (Phosterodoros) jonesi Stone and Snoddy..
SimuZium (Phosterodoros) lake~i Snoddy.. .....
SimuZium (Phoster~odoros) notiale Stone and Snoddy.
SimuZium (Phosterodoros) nyssa Stone and Snoddy..
SimuZium (Phosterodoros) taxodium Snoddy and
Beshea-r. . . . . . . . .
Simu*Zium (Psilozia) vistatum Ze~ttrstedt
Simulium (Simutium) decorun Walker ........
Simulium (SimuZium) tuberosum (Lundstram)......
SimuZium (S~imulium) verecundwnr Stone and Jamnback..
Opephia species Undetermined No. 1.. ......
Shaulium species Undetermined No. 1.... ....

Leucocytozoon smith Transmission... ........

V. CONCLUSIONS.. ... .. .......

LITERATURE CITED. ...... ........

APPENDIX. COLLECTION SITE NAMES AND LOCATIONS........ .

BIOGRAPHICAL SKETCH.. ... .............
















LIST OF TABLES


>le Pag~e

1. Florida black fly species. .. .. .. .. .. ... 55

2. Florida black fly distribution records by county .. .. 57

3. Black fly associations based on! collections of immature
stages .. .. .. ... .. ... .. .. ... 59

4. Black flies captured in Manitoba traps . ... .. .. 60

5. Sentinel turkey locations and results. .. .. .. .. 278

6. Black flies captured in ramp traps .. .. .. .. .. 281

7. Blackout box trapping results. . .. ... .. ... 282

8. Black fly captures from exposed turkeys. .. .. .. .. 283

9. Leucocytozoon smith transmissions .. .. . ... 289
















LIST OF FIGURES


Igure Page

1. A modified Manitoba trap with a black plastic skirt . .. 44

2. Sentinel turkeys in an exposure cage. .. .. .. .. .. 47

3. A blackout box trap in the field. .. . .. ... ... 47

4. One view of a ramp trap .. .. .. ... ... .. 48

5. An exposed turkey in the field. .. .. ... .. .. 48

6. Glass container and paper cartons used for holding
black fly adults alive in the laboratory. .. .. .. .. 51

7. Locations in Florida where black flies have been
collected . .. .. . .. . .. .. .. 56

8. Seasonal occurrence of black flies in Florida .. ... 63

9. Dorsum of the head capsule of a black fly larva
(S. slossonae) .. .. .... .. .. .. .. .. 65

10. Venter of the head capsule of a black fly larva
(C.ornithoph~iia ... .. .. . .. .... .. .. 65

11. Lateral view of two black fly larvae (S. dirEense) .. .. 66

12. Black fly pupa and cocoon (S. diriense). .. .. . ... 66

13. A wing of the black fly onephia ornith~ophilia .. .. .. 68

14. A frontal view of the head of a female black fly
(S. notiate). .. .. .... .. .. . ... . .. 68

15. The male genitalia of a black fly, Cnephia
ornithoohilia ... .. .. .. . ... ... . .. 69

16. The distal portion of the hind leg of a S. meridionate
female. .. ... .. ... .. .. .. . .... 69

17. The terminalia of a female S. merZidonatea .. .. .. .. 70

18. The head spots of a larva of C. ornithophiflia . ... .. 82


vii










LIST OF FIGURES
(Continued)


Figure rage

19. The pupal exuvium and cocoon of C. ornithophilia. . ... 82

20. Tarsal claw of a female of C. ornithophilia .. .. .. 84

21. Genital fork and terminalia of a C. ornithophilia female. 84

22. Collection locations for C. ornithophilia in Florida. .. 88

23. Site 119, Guim Creek, a stream inhabited by
C. ornith~ophilia. .. . ... .. . . .89

24. The pupa and cocoon of S. mer~idionate .. ... . 94

25. The scutum of' a S. meridionale female . .. .. 94

26. Collection locations for S. meridionale in Florida. .. 97

27. Gular notch of a S. sZossoncae larva .. .. .. .. .. 101

28. The pupa and cocoon of S. slossonae . ... .. .. .. 101

29. Terminalia of a male of S. s~ossonae. .. .. .. .. .. 102

30. Terminalia of a female of S. sZossonae. .. .. .. .. 102

31. Collection locations for S. slossonzae in Florida. .. .. 105

3.Site 43, Double Run Creek, where S. so~;ssonae
immatures were collected. . .. .. .. .. .. 107

33. Cephalic apotome of a S. congareenarum larva. . .. ... 118

34. Venter of the larval head capsule of S. congareenaumo .. 118

35. Pupa and cocoon of S. congareenarum .. .. ... .. 120

36. Terminalia of a male of S. congareenarum. .. .. ... 120

37. Terminalia of a female of S. congaree~nawn. .. .. .. 121

38. Collection locations for S. congereencrum in Florida. .. 124

39. Site 216, Turkey Creek, a typical S. congareenarum
stream. .. .. ... .. .. .. .. .. .. ,.. 125

40. Dorsal view of the head capsule of a S. didcense larva. . 130

41. Gular notch and by~postomiumn of a S. d'ixiense larva. . .. 130









LIST OF FIGURES
(Continued)


vFigure.

42.

43.

44.

45.



46.

47.

48.

49.

50.


.. .132

. 132

. 134


Male terminalia of S. diziense. ..........

Terminalia of a female of S. dizciense .......

Collection locations for S. dizciense in Florida ..

Site 74, Pine Barrens Creek, a stream inhabited by
S. didcense . . . . . . . . . .


Cephalic apotome of a S. haysi larva. .......

Gular notch of a S. haysi larva ..........

Pupal exuvium and cocoon of S. haysi. .......

Location of the collection site for S. hacysi in Flol

Site 195, Juniper Creek, where S. haysi was collect~


. .135

. .138

..138

. .139

. .141

. .142


rida

ed .


51. Cephalic apotome of a S. jenningsi larva. .. .. ... 145

52. Gular notch of a S. jenningsi larva . ... .. .. .. 145

53. A pupa and cocoon of S. jenningsi .. .. .. .. .. .. 146

54. S. jenningsi male terminalia. .. ... .. .. .. .. 146

55. Genitalia of a female of S. jenningsi .. .. .. . 147

56. Collection locations for S. jenn~ingsi in Florida. . ... 151

57. Site 141 at Gulf Hamrmock where S. jennings~i was collected 152

58. Cephalic apotome of a S. jonesi larva ... .. .. 156

59. Gular notch of a S. jonesi larva. .. .. .. . .. 156

60. Respiratory organ of a S. jonesi pupa .. .. .. . 157

61. Terminalia of a male of S. jonesi .... .. .. .. 159

62. Genital fork and terminalia of a female of S. jon~esi. .. 159

63. Florida collection locations for S. jonesi. .. .. .. 161

64. Site 210, the Senholloway River, a S. jcnesi collection
location. ... .. .. .. .. .. .. .. .. .. 162










LIST OF FIGURES
(Continued)


Figure

65.

66.


67.

68.


69.

70.


71.


72.

73.

74.

75.


76.

77.


78.

79.


Page

171

171

172


172

. .173


. .173

175


. .176

. .183


...183

. .184

184

185


. .185

. .187


Cephalic apotome of a S. lakei larva. ........

Gular notch and hypostomium of a S. lakei larva ...


Pupa and cocoon of S. lakei .............

Dorsal view of a male of S. laket .....

Terminalia of a male of S. lakei....... .. .


Terminalia of a female of S. lakei. .........

Collection locations for S. lakei in Florida. ....


Site 135, Otter Creek, a collection site for S. lakei


Head spots of a S. notiale larva. ..........

Gular notch of a S. notiate larva ..........


Pupal exuvium and cocoon of S. notiale. .......

Scutum of a male of S. notiate. ...........


Terminalia of a male of S. notiale. .........

Terminalia of a female of S. notiaie. ........

Collection locations for S. notCate in Florida. ...


80. Site 88 at Chattahoochee where S. notiate immatures
were found. .. ... .. .. .. . .. .. . 188


81. S. nyssa pupa and cocoon. .. ... .. ... .. .. 191


82. Collection locations for S. nyssa in Florida. ... .. 193


83. Site 116, Blue Creek, where S. nyssa was collected. . .. 194

84. Cephalic apotome and head spots of a S. taxcodium larva. . 197

85. Gular notch of a S. tarcodium: larva. .. ... .. .. 197


86. Pupal exuvium and cocoon of S. taxcodiu7 . ... .. .. 198

87. Male terminalia of S. txodium. . .. .. .. ... .. 198


88. Female terminalia of S'. Laod~ium. .. .. .. .. .. .. 199




































































110. Terminalia of a female of S. tCuberosumr. .. ... . .. 239

111. Collection locations for S. tubrerosum in Florida. .. .. 243

112. Site 56 in Clay Countyr, a collection spot for S. tuberosum. 266


LIST OF FIGURtES
(Continued)


site. ...


Page

201

203

209

209

210

210

211

211

217

219

226

226

227

227

228

232

234

237

237

238

238


89. Collection locations for S. tax~odium in Florida

90. The Ichetucknee River, a S. taxcodium collection

91. Head spots of a S. vittatum larva .......


92.

93.

94.

95.-

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

107.

108.

109.


Gular notch of a S. vittatum larva. ............

S. vittatum pupa and cocoon ................

Terminalia of a S. v-ittatum male. .............

Scutum of a female of S. vittatum.... .....

Terminalia of a female of S. vittatum... ....

Collection locations for S. vittatum1 in Florida ......

Site 167 at Crestview where S. vittatum was collected ...

Head spots of a S. decorum larva. .............

Gular notch of a S. deconrum larva .............

S. decorum pupa and cacoon. ................

Male terminalia of S. decoum ..... .....

Female terminalia of S. decorum ..............

Collection locations for S. decorum in Florida. ......

Shepard's Mill, Site 86, where S. decorum was collected ..

The cephalic apotome of a S. tuberosum larva. .......

The gular notch of a S. tuberosum larva ..........

Pupa and cocoon of S. tuberosum.... .....

Male terminalia of S. tuberosum.... .....









LIST OF FIGURES
(Continued)



Figure Page

113. The head spots of a S. verecundum larva. .. .. .. .. .257

114. The gular notch of a S. vereoundum larva .. . .. ... .257

115. The pupa and cocoon of S. verecundum .. . .. .. .258

116. Male terminalia of S. verecundum .. .. .. .. .258

117. Terminalia of a female of S. vereoundum. .. .. .. .. .259

118. Collection locations for S. verecundum in Florida. . .. .262

119. Panther Creek, Site 223, where S. verecundumlu was collected .264

120. Cephalic apotomce of a larva of Onephia species No. 1 .. .269

121. Venter of the head capsule of a larva of Cnephia species
No. 1... . .. .. .. .. . . .. .. .. 269

122. Collection location for Q~nephia species No. 1 in Florida .270

123. Pupal exuviumn and cocoon of SimuZi~um species No. 1 .. .273

124. Larval head spots of Simulium species No. 1. .. .. .. .274

125. Venter of the head capsule of the larva of SimuZium species
No. 1. .. .. .. .. .. .. .. ... .. ... .274

126. Collection location for SimuL~ium species No. 1 in Florida. .275

127. Small flow to Pine Barrens Creek, Site 74, where
Simulium species No. I was collected .. .. . .276

128. Gametocytes (G) of L. smith among normal turkey blood
cells. .. .. .. .. . . . .. .. .291

129. 00kinetes of L. smith .. .. .. . . .291

130. Sporozoites of L.. smith photographed in saline. . ... .292

131. Stained sporozoites of L. smithi . . .. .. . .292















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 BLACK FLIES (DIPTERA: SIMULIIDAE) OF F~LORIDA AZND THEIR
INVOLVEMENT IN THIE TRANSMISSION OF
Leucocytozoon smith TO TURKEYS

By

Dennis Drew Pinkovsky

August, 1976


Chairman: Jerry F. Butler
Major Department: Entomology and Nematology

Immature and adult black flies (Diptera: Simuliidae) were collected

in Florida over a period of three years. Eighteen species of black

flies including ten which are new records for the State were found to

occur in Florida. Records of black flies from 192 locations in 50

counties are included. Biological information is provided for each

species together with data on distribution, seasonal occurrence, scream

ecology and species associations. Keys to the Simuliidae of Florida are

provided and structures are illustrated. Representative specimens have

been deposited in the Florida State Collection of Arthropods and in the

United States National Miuseum, Washington D.C.

Transmission investigations have incriminated three species,

SimuZiumi congareenrum~, S. me~ridion~ale, and S. slossonae, as vectors of

Leucoceytozon smithi to turkeys in Florida. On nineteen occasions

L. smithli was transmitted to domestic turkeys by the bites of infected

black flies.


xtiii














CHAPTER I
INTRODUCTION



Members of the family Simuliidae as immatures inhabit flowing water

and as adults are often blood feeders which may vector diseases to man

and animals in many parts of the world. A knowledge of the composition,

distribution, ecology, and habits of the simuliid fauna of any area of

local concern is essential for proper assessment of the impact on man of

these insects. Successful black fly control is dependent on thorough

knowledge of the major breeding sites, seasonal occurrence and other

facts concerning local Simuliidae.

When I arrived in Florida to begin my Ph.D. research I wrote to

the Florida Division of Health offices in Jacksonville, Vero Beach, and

Panama City and enquired about previous investigations on the black

flies of Florida. I found that little work had been done in the State

on this family of biting flies (Beck, 1973; Linley, 1973; Rogers, 1973 -

all personal communications). Mrs. A.T. Slosson collected black flies

in Florida (Dyar and Shannon, 1927), probably at the turn of the

twentieth century. Some of Mrs. Slosson's specimens and a portion of

those of Calvin Jones, Harry Couck, Darrell Anthony and other USDA

researchers who collected black flies in the northern and central por-

tions of the State in the 1940's and 1950's are located in the U.S.

National Museum. Based at least in part on the examination of

these specimens Stone (.1965.) listed the following seven species from

Florida: Crnephia (Cnephia) pecuarum (Riley); Simulium (Eusimuli~un)









congareenatnum(Dyar and Shannon); Simulium (Byssodon) meridionaite Riley;

Simuliwn, (Byssodon) saossonae Dyar and Shannon; Simulium (Simutium)

decorum Walker; SimuZitrm (Simulium) jenningsi Malloch; and Sirmutium

(Simutium) tuberosum (Lundstrijm). Stone and Snoddy (1969) did not list

S. jenningsi as occurring in Florida but did record two new species

Simulium (Phosterodoros) jonesi Stone and Snaddy and Simulium (Phostero-

d'oros) nyssa Stone and Snoddy from the State. Thus the total number

of species of black flies known to occur in Florida when I began my

research was eight.

This research was initiated in the fall of 1973 with the following

objectives: 1) to determine the species complement, distribution and

seasonal occurrence of black flies throughout Florida; 2) to gather

ecological information primarily on the immature stages; 3) to provide

keys with illustrations to the black flies of Florida; 4) to gather and

deposit in the U.S. National Museum of Natural History and the Florida

State Collection of Arthropods as complete a collection of adult and

immature Florida black flies as possible; and 5) to determine the vec-

tors of Leucocytozoon smith in turkeys in Florida.













CHAPTER II
LITERATURE REVIEW



Simuliidae


Taxonomy and Distribution


Rubtsov (1974) presents a concise discussion on the history and

major advances in black fly taxonomy. Linnaeus (1758) first described

two simuliids and assigned the names CuZez reptan~s and Culex: equinzus

but did not differentiate them from mosquitoes (Davies et al., 1962).

Latreille created the name Simulium for the genus in 1802 using Rhagio

cotomb~aschenzsis Fabricus as the type (Stone and Jamnback, 1955). This

remained the only genus for the thirty to forty black fly species de-

scribed up to the turn of the twentieth century. The work of Smith and

Kilbourne (1893) and others which focused on arthropods as vectors of

diseases stimulated much research on insects and related groups of

medical and veterinary importance. Roubaud (1906) designated two black

fly subgenera, Prosimuium and Eusinulium, based on differences in wing

venation. Lundstram (1911) in Sweden and Jobbins-Pomeroy (1916) in the

United States were early taxonomists who made use of the male terminalia

of simuliids to separate species. Edwards (1915) in England also used

male terminalia but more significantly in 1920 stressed the importance

of studying black fly larvae and pupae. Dyar and Shannon (1927) first

used female genitalia to separate North American black flies. Enderlein

(1930) divided the family Simuliidae into 29 genera. Smart (1945) took

a more conservative approach and distinguished 6 genera in the family:








ParasimuZium Malloch, ProsirmuLium Roubaud, Aulstrosimulium Tonnoir,

Gigantodcw: Enderlein, Onephia Edwards and Simulium Latreille. Stone

(1963) listed 11 genera and 22 subgenera in the Simuliidae which he

considered valid. Crosskey (1969) stressed a trinomial approach to

simuliid taxonomy with heavy reliance on subgenera. Rubtsov (1974)

recognized 17 genera of black flies in the Palaearctic region and listed

59 genera for the Simuliidae of the world, many of which American and

British authors regard as subgenera.

Brues et al. (1954) provide a worldwide bibliography of important

papers which deal with the taxonomy of the Simuliidae up to the early

1950's. In addition to the works already mentioned outstanding publi-

cations since about 1950 dealing with the classification and distribu-

tion of black flies outside of the United States include:


Palaearctic region Davies (1968) Britain, Carlsson (1969) Spain,

Rubtsov (1956 and 1962) U.S.S.R., Kuusela (1971) Finland,

Zivkovic (1971) Yugoslavia, Rivosecchi (1971 and 1972) Italy,

Crosskey and Pete~rson (1972), and Zwick (1974) Germany;

Ethiopian region Crosskey (1957) West Africa, Travis et al. (1967),

Crosskey (1969) Africa and its islands, Fain and Elsen (1973) -

Cameroons, Lewis and Raybould (1974) Tanzania;

Oriental region Travis and Labadan (1967a) Asia and European U.S.S.R.,

Delfinado (1969, 1971) Philippines, Crosskey (1973), Takaoka

(1973) Nansei Is., Uemoto et al. (1973) Japan, Lewis (1973) -

Pakistan, Datta (1973, 1974, 1975) India;

Australian Dumbleton (1963, 1972) Australia and New Zealand, Travis

et al. (1968);

Neotropical Leon and Wygodzinsky (1953) Ecuador, Dalmat (1955) -









Guatemala, Vulcano (1967), Travis and Labadan (1967b),Barreto

(1969) Colombia, Wygodzinsky and Coscaron (1970, 1973), Wygod-

zinsky and Najera (1970), Wygodzinsky (1971) Northern Andes,

Perez (1971) Venezuela, Rubtsov and Avila (1972) Cuba, Travis

et al. (1974);

Nearctic (excluding the U.S.) -- Syme and Davies (1958), Davies et al.

(1962) Ontario, Wood et al. (1963) Ontario, Travis et al. (1969),

Peterson (1970) Canada and Alaska, Lewis and Bennett (1973) -

Newfoundland.


In addition to the above works those of Rubtsov (1970, 1974) and

Coscaron and Wygodzinsky (1973) mention the variability which has been

observed in characters that are used for taxonomic determination of

black flies. Gambarian and Terterian (1973) applied a numerical tax-

onomy approach and using 100 characters tried to separate simuliids in

the Eusimulium group. The subgenus was found to be very homogeneous

and creation of supraspecific taxa was not statistically justified.

Internationally, black fly species determinations based on classical

external morphological characters and behavioral differences are being

supported by cytological, specifically chromosomal, studies. Rothfels

(1956) provided an introduction and overview on analytical techniques

for cytologically comparing black flies. Landau (1962), Dunbar (1969),

and Vajime and Dunbar (1975) used chromosomal differences to separate

forms or sibling species of a variety of simuliids including disease

vectors where previously each had been considered a single species.

Taxonomic works covering the whole United States include: Coquillett

(1898, 1902), Malloch (1914), Jobbins-Pomeroy (1916), Dyar and Shannon

(19217), and Stone (1965). Other publications were more regional in








scope. Western and midwestern articles include: RTinn (1938), Smith

and Lowe (1948) California; Nicholson and M~ickel (1950) Minnesota;

Stone (1952), Somrmerman (1953) Alaska; Wirth and Stone (1956) -

California; Stone and DeFoliart (1959); Anderson and Dicke (1960) -

Wisconsin; Stone and Boreham (1965), Hall (1972, 1974) California;

and Corredor (1975) Washington.

Northeastern U.S. works include: Johannsen (1903), Leonard (1926),

Metcalf (1932), DeFoliart (1951), Stone and Jamnback (1955), Jamnback

and Stone (1957), Jamnback (1969), Pinkovsky (1970), and, reporting the

unusual find of a South American black fly in the U.S. Wygodzinsky (1973)

- New York; O'Kane (1926) New Hampshire; Frost (1949) Pennsylvania;

Dimond and Hart (1953) Rhode Is.; Sutherland and Darsie (1960a and b)

- Delaware; Stone (1964) Connecticut; Holbrook (1967) Massachussetts;

Eckhart and Snetsinger (1969) Pennsylvania; and Crans and McCuiston

(1970a) New Jersey.

In the southeastern U.S. Tucker (1920) gave accounts of personal

experiences with black flies and reported two species, S. pecuarun and

S. meridionale, from Louisiana. Jones and Richey (1956) conducted a

survey of the black flies of Jasper County, South Carolina, and discussed

the biology, ecology and relationships to Leucocytozoon in turkeys of

one Cnephia (pecuarum) and seven Simulium species (congareenarum, deconk ,

jenningsi, slossonae, tuberosum, voenuatum and undescribed species).

Snow et al. (1958) reported on the ecology, habits and distribution of

black flies occurring in the Tennessee River Basin and recorded species

in the three genera: Chephia mutate2), Prosimutium~ (hirtipes, magnum,

plus undescribed species), and Silmu~ium (decorum, fibrinflatum~, jenningsi

ap. group, merid~ionate, pictipes, tuberose, veuan~tum, verecundum,





-7-


vittatum and three undescribed species). Snoddy and Hays (1966) mentioned

that eleven species of Simuliidae were captured at one location in

Alabama using a New Jersey light trap modified for daylight use by

removing the light and substituting,as an attractant, carbon dioxide

gas dispensed at .45 kg (1 lb)/hr. Stone and Snoddy (1969) present

distribution records, descriptions, and some biological information for

28 species of black flies discovered or expected to occur in Alabama.

Garris et al. (1975) present the seasonal distribution of 7 species of

Simulium collected from streams in Sumter Co., South Carolina, and

mention observations of black flies feeding on turkeys and captured in

a CO2-baited trap. Snoddy and Beshear (1968), and Snoddy (1971, 1976)

describe and give some facts on the biology of three new species

(S. tcaodiumi, podocstem~i and Zakei) in the expanding species list of the

former S. jenningsi group.



Bionomics


The eggs of black flies are dropped freely into the water or are

attached to substrates in the flows of streams and rivers. The larvae

which develop feed by filtering material from the current or by scraping

organic matter off the substrate and, usually within a month, transform

into pupae. From the pupa which is normally attached to rocks or

vegetation beneath the surface of the stream an adult fly emerges, in a

bubble of gas, rises to the surface and is immediately capable of flight.

The adult females of most black fly species suck blood and both male

and female flies obtain energy from natural sugar sources such as nectar

and honeydew.

Eggs. Ussova (1961) describes most black fly eggs as between .24








and .33 mm long and irregularly triangular in shape with rounded corners;

the eggs of some Cnephia species, however, are said to be larger and

elongate elliptical. Golini and Davies (1975) found black fly eggs were

.228 mm long and .139 mm wide. Davies and Peterson (1956) examined the

eggs of four genera (Gymnopais, ProsimuZium, Cnephia, Simutium) and

found no external sculpturing. The eggs are light in color when just

laid and darken as the embryos develop. Davies and Peterson (1956)

found that the eggs of Gymnopais and Prosimulium were the largest and

those of ProsimuZium and C~ep~hia were the narrowest of the genera checked.

Black fly eggs are often found attached to vegetation and rocks in a

swift flowing stream or river or lying free on the stream bottom.

Tarshis (1968) conducted experiments on and reviewed accounts in the

literature of desiccation and the overwintering of black fly eggs and

concluded, as Wu (1930) had done earlier, that only those eggs which

are kept moist, by damp stones or underground springs, etc., in ap-

parently dry stream bottoms and other situations are able to remain

viable. Tarshis (1968) found that freezing eggs of a number of species

at 0 to-700C killed the embryos, while he was able to maintain moistened

eggs alive for 424 days at 2-90C. Kurtak (1974) found that eggs of

SimuZ~ium pictipes, recovered while encased in ice from rock crevices

along a stream, hatched in the lab after three days in 100C flowing

water. In northern areas eggs of many black fly species hatch when the

water temperature reaches around 80C (Carlsson, 1967). Raybould and

Grunewald (1975) found eggs of the K~ibwezi form of S. down2osrum watch at

200C four days after oviposition. Tarshis (1968) indicates normal egg

hatch for Maryland black fly eggs occurs within one to five days.

Larvae general morph~ology and development Black fly larvae are

usually between 4.5 andl 10 mm long when mature, club-shaped with a wider









posterior end, and possess a pair of multiple-rayed cephalic fans on

the anterior end, an unsegmented ventral proleg in the thoracic region,

and a circlet of anal hooks posteriorly. Dark cephalic head spots mark

the origins of cephalic muscles. Tarshis (1968) found that larvae

hatched from eggs in the lab after 7-38 days at 1000 and 1-5 days at

20-250C. Larvae developed to pupae in 18-50 days at 100C and 11-29 days

at 20-250C. Cameron (1922), Dalmat (1955) and Reisen (1975) found six

larval instars in the black flies they studied. Johnson and Pengelly

(1970) observed S. rugglesi to pass through seven larval instars and

Fredeen (1975) mentioned a seventh and final instar for S. aration.

Craig (1975) distinguished nine larval instars in Tahitian species of

black flies.

Larval habitat. Larvae frequently attach by their posterior hooks

onto a patch of silk secreted by their large, looped salivary glands.

The silk is often applied to the same substrate on which the eggs are

found. Larvae may drift a short distance tethered to the old substrate

by a silk thread (Tarshis and Neil, 1970). A larva may also move in a

looping, geometrid fashion from silk patch to silk patch alternately

attaching its proleg then its anal disc (Dalmat, 1955). Elliot (1971)

reported that black fly larvae can actively migrate upstream as well as

passively downstream. Clean, smooth items free of algae or slime

are preferred attachment substrates for the larvae and larvae are

often located on such objects in stream sections where the current

is increased by a partial obstruction (Dalmat, 1955; Carlsson, 1967).

Larvae show positive phototaxis and colonize light substrates faster

and more densely than dark substrates (Carlsson, 1967). Unusual

attachment sites include the phoretic associations of black fly larvae

on prawns, mayfly nymphs, crabs, and other arthopods reported by









Disney (1971, 1973, 1975) and thle attachment of young larvae of S.

damnosum to older larvae of the same species (Burton, 1971). Carlsson

(1967) in Sweden found the greatest concentrations of larvae in flows

80 to 120 cm /sec although some species preferred 40

cm/sec currents. Cariase (1962) in the Philippines found immature black

flies concentrated where the water velocity was .5-.86 m/sec (1.63-2.83

ft/sec). Wu (1930) reported that simulfid larvae remained well estab-

lished at a velocity of 1.83 m/sec (6 ft/sec). Rohdendorf (1974) sug-

gested that the closed respiratory system of black fly larvae resulted

after invasion of the swift flowing well oxygenated habitat typical of

most simuliids. Tarshis (1968) found that black fly larvae could not

live more than eight hours in still water. Anderson and Shemanchuk (1975),

however, report they transported black fly larvae for several days in

shallow, non-agitated, ice-chilled water with little mortality. Wu

(1930) found approximately equal amounts of dissolved oxygen in quiet

and turbulent sections of black fly streams and after a number of ex-

periments concluded that black fly larvae have a definite requirement

for current to lessen sedimentation and supply adequate food, not because

of improved oxygen conditions. Crans and McCuiston (1970b)found larvae

Ln permanent rivers and streams, temporary creeks and flowing roadside

ditches. Dalmat (1955) and Lewis and Bennett (1975) reported finding a

few live larvae in situations where the flow was so slow that silt and

mud present prevented the larvae from anchoring to any fixed object.

Van Someren (1944) reported S. ruficorne larvae from small pools and

footprints in sandy river beds in Somaliland. Certain species are

typically found in limited type's of lotic habitats: S. arcticum in

large rivers (Cameron, 1922), S. pictipes at the outflow of damns and on









flat bedrock (Snow et al., 1958) and S. cohraceum in very small streams

about one meter (a few feet) wide and a few centimeters (a few inches)

deep (Dalmat, 1955). Travis and Vargas (1970) found that in contrast

to the clear mountain streams, the lower, slower Costa Rican streams

most polluted with sewage and garbage had the greatest larval and pupal

populations. Cariaso (1962) reported that black fly larvaeweredis-

couraged from breeding in water polluted with nitrogenous and human

wastes. Carlsson (1967) indicates moderate pollution increases organic

drift and is good for larvae but heavy pollution clogs the cephalic

fans. Streams in Wisconsin carrying large amounts of eroded soil par-

ticles and other detritus were observed to be poor habitats for most

black fly larvae due to feeding interference (Anderson and Dicke, 1960).

Larvae feeding structures, behavior, and food. Davies (1974)

and Craig (1974) describe the evolution and development of the important

larval feeding structures in many species, the cephalic fans, and men-

tion how these lateral palatal brushes which are well developed filter-

ing devices in most black flies are apomorphically absent in later

instars of Gyrmnopais and Zainnia and modified into raking structures

in Crosetia species. Chance (1970) stated that particles from about 1

to 350 microns in diameter, the majority in the 10 to 100 micron range,

were ingested by four filter feeding species she studied. Larvae were

found to attach within 10 centimeters of the water surface, extend their

bodies into the flow and alternately open and close the fans combing

particles from the fans when they were closed into the cibarium where

a bolus was formed. Mulla and Lacey (1976) report larvae, with their

heads pointed downstream, attach to silk deposits with their anal hooks,

rotate the body 900 to 1800 and open the cephalic fans to strain matter









from the flow. Chance (19703) suggests that the horizontally operating

mandibles, especially in non-filtering, grazing forms like Tannia,

scrape organic food off the substrate. Rubtsov (1974) discusses the

outer sclerite of the labium the submentum or hypostomum, an anteriorly

serrate structure bearing setae which lies just cephalad of the

gular notch and mentions that it is a tactile organ, is important in

producing the larval silk strands, and that it may serve to scrape food

from the substrate. In species which lack fans the submental teeth

sometimes acquire a spatulate shape.

Jobbins-Pomeroy (1916) reported Euglenaand Spirogyra were important

food items for black fly larvae but Cameron (1922) found diatoms formed

the main components of the food. Anderson and Dicke (1960) found

diatoms, other algae, and considerable inorganic material in the in-

testinal contents of black fly larvae. Carlsson (1967) listed as larval

food bacteria, plankton, plant and animal parts, pollen and aerial

fallout. Spring flooding leads to large outbreaks of black flies when

rising water temperatures trigger synchronous hatching of many black

fly eggs. The rising stream and river waters make more substrate avail-

able for larval attachment and increase organic debris and hence food

available for the larvae. Bacteria filtered out by S. underhilli led

to speculation of using simuliid larvae as indicators of pollution

(Snoddy and Chipley, 1971). Reisen (1974) found that larvae remove

bacteria and organic and inorganic particulate matter. Young larvae of

Simulium species in nature fed at a faster rate than did older larvae

as indicated by the time necessary to void a plug of dye particles

(Mulla and Lacey, 1976). These authors also found that at a lower

temperature (12.80C 550F) a longer time (35-55 min) was necessary to









eliminate a particulate plug from larval guts than at a higher tempera-

ture (300C = 860F, 20-30 min).

Pupae. Pupae attach to the same substrates as the larvae. The

larvae form silken cocoons in which pupation occurs. Hinton (1958)

mentions a pharate pupa stage where pupal characters are visible within

the last larval skin but the larval mouth parts are still articulated

and feeding continues up to when this "pupa" begins spinning the cocoon.

The cocoons may be thin and rudimentary (Rrinnia), shapeless, irregular

masses of silk (Prosim~utium and some Cnephia), sturdy, coarse cocoons

(Sinuium pictipes) or tapered, slipper-like finely shaped and tightly

woven cocoons (most SimuZium) (Stone and Jamnbackc, 1955). Members of

the Simuliumn subgenus Phosterodoros have cocoons with forward edges that

are convex in profile and have a large opening anteriorly on each side

of the cocoon (Stone and Snoddy, 1969). Field and Law (1961) describe

what Underhill (1944) illustrated for S. jenningsi (as S. nigroparvum),

that is, sexual dimorphism in the cephalic plate of simuliid pupae. The

cephalic plate, a sclerite that begins between the bases of the antennal

sheaths and extends over the cephalic area, is longer and narrower

in the male than it is in the female pupa. The pupae usually face down-

stream and are bordered on each side by tubular filaments which vary in

number and shape according to species and aid in respiration. Cameron

(1922) observed that the number of pupal filaments in S. arcticumt varied

from the usual 12 to 13 or 11 and that one set might vary while the

other group on the same pupa might bear the more typical number of

filaments. Coscaron and Wygodzinsky (1973) mention the variation in

the point of bifurcation or petiole lengths of the respiratory filaments

of different specimens within the same species.









Adults emergence. Tarshis (1968) reported that pupal develop-

ment of three species of Simulium took 1 to 5 days at 20-250C and that

at 100C it took 21 days following pupation before adults began emerging.

Davies et al. (1962) describe how, at emergence, after gas fills the

skin of the submerged pupa, the adult ruptures the pupal exuvium, pulls

itself through a T-shaped opening and quickly floats to the surface in

a bubble of gas (also see Hannay and Bond, 1971, section below). Carlsson

(1967) found that a heavy silt cover on pupae prevented adults from

emerging. Adults may easily be reared from pupae placed on moist filter

paper in petri dishes (Hannay and Bond, 1971; Sutcliffe and McIver,

1974). Wenk (1965) found males emerge sooner from pupae than do females.

Disney (1969) found three species of Simulium in Africa pupate by day

and may prolong pupation to avoid emerging at night. On warm days

eclosion occurred early in the morning, on cooler days eclosion peaks

occurred during the late morning and mid-day and under artificially cold

days emergence was shifted to the late afternoon.

Adults appearance and morphology. Adult black flies rarely ex-

ceed 5 mm in length (Dalmat, 1955). Male and female black flies are

hump-backed in appearance. The males are holoptic, usuallydarkerand

morevelvety than females and often bear a shiny pair of anterior lateral

spots on the scutum (Davies et al., 1962). Costalization (i.e., a

strengthening of the anterior wing veins and a decrease in wing venation

in the posterior portion of the wing), shortening of the legs, evolution

of the calcipala and pedisulcus, increase in head size and shortening of

the abdomen with reduction of tergites and sternites are considered

apomorphic developments (Rohdendorf, 1974; Rubtsov, 1974). Hungerford

(1914) in discussing the anatomy of Simulium vittatumt adults noted that









black fly females possess a single spermatheca. Lewis (1957) presented

morphological information about adults of S. damnnsum and suggestions

on dissection techniques. Bennett (1963b)found the shape and appearance

of the salivary glands of adults valuable in differentiating species.

Hannay and Bond (1971) found raised cylindrical buttons, each with a

wax filament, between the macratrichia on black fly wings and suggest

these structures may aid the adult in keeping its wings dry during

emergence. Sutcliffe and Melver (1974) describedcleaning hairs and

combs of cuticular teeth on the metathoracic legs and cleaning hairs

on the prothoracic legs which are used to clean the wings and head

appendages, respectively.

Adults attraction and trapping. Wirth and Stone (1956) indicate

that male black flies are readily collected at light traps. Snow et al.

(1958) captured adults at light, on vegetation, in cars and biting

mammalian hosts. Fallis and Smith (1964) succeeded in using ether

extracts of birds as attractants for simuliids. Anderson and DeFoliart

(1961) used a variety of caged birds to attract ornithophilic black

flies to traps in Wisconsin. Golini and Davies (1971) found that fe-

males of S. venustum fly upwind to a carbon dioxide source and cease

flying upwind if the CO2 source is turned off. Snoddy and Hays (1966)

and DeFoliart and Morris (1967) made use of the attractancy of CO2! to

capture simuliids. The silhouette of a target rather than the reflec-

tance is more important in attracting black flies (Gillies, 1974).

Peschken and Thorsteinson (1965) state that black flies are more at-

tracted to stationary targets than to moving targets and simuliids show

less discrimination between form and shape of three dimensional objects

than some other hematophagous insects. Bradbury and Bennett (1974a)






-16-


found bloodseeking black flies in the genera Prosimulium, Onephia, and

Simuliwn were more attracted to black, blue, and red than white or

yellow, especially when the colors were of low visible reflectance.

Thorsteinson et al. (1965) describe a tripiod canopy-type Manitoba fly

trap which uses a large black sphere target to make the trap visually

attractive to bloodsucking diptera. Bradbury and Bennett (1974b) found

that black flies could distinguish between targets based on their color,

largely independent of carbon dioxide flow rates. Carlsson (1967) stated

lakes, shoals, bogs and swamps act as "collecting mirrors" for female

black flies seeking ovipositioning spots. Female black flies seeking

oviposition sites oviposited more on green and yellow colored plastic

strips in a stream than on red, purple, white or black strips (Golini

and Davies, 1975). The Malaise trap, a final valuable collecting device

for black flies and other insects,was conceived as a passive or random

flight interception trap;however there is some evidence that the degree

of color contrast between the trap and background vegetation is an

attractance factor (Roberts, 1970 and 1972).

Adult mating. Females of C. de~cotensis have mature eggs at the

end of the pupal stage and copulate with males on damp rocks soon after

emergence. In general, however, simuliids can mate any time from

emergence to oviposition time (Davies and Peterson, 1956). Davies and

Peterson (1956) were unable to induce male black flies to mate by broad-

casting the sound of the female wing beat frequency. Compared to males

which do not form mating swarms, males with mating flights have larger

eyes and enlarged upper facets. Snow et al. (1958) observed that males

of S. pictipes flew upside down with their abdomens curved upward, con-

tacted females as they flew by overhead, settled to a solid substrate,









and copulated venter to venter with the female situated uppermost. The

presence of about one male for every two hundred females of onephia mutata

is considered evidence for parthenogenic reproduction (Davies and Peterson,

1956). Downes (1965) reports Prosimulium .ursinum is a maleless species

in which the females do not even emerge from the pupa but disintegrate

shedding the eggs into the stream.

Adult feeding. Black flies have been collected from flowers

(Davies and Peterson, 1956). Hocking (1953) stated that black flies and

other northern biting flies obtain the energy for flight almost entirely

from floral nectar. Lewis and Domoney (1966) reviewed the importance of

sugar feeding on bloodsucking, autogeny, and parasite development and re-

ported finding glucose, sucrose, fructose and other sugars in the crops

of 101 wild caught Simutium. There are scattered reports of black flies

being attracted to and feeding on cold-blooded animals (Hagen, 1883;

Jobbins-Pomeroy, 1916; Smith, 1969), but of more importance are their

ornithophilic, mammalophilic, and anthropophilic bloodsucking habits.

Onephia dacotensis, Gyrmnopais hotoptious, Tw~innia ti~bbesi and a few

other black fly species acquire sufficient nutrients during the larval

stage to produce eggs and do not suck blood (Davies and Peterson, 1956;

Shewell, 1957). Rohdendorf (1974) suggested that limited larval food

stimulated the adult female black flies to hunt and feed on vertebrates.

Cameron (1922) mentions the tenacity and persistence black flies ex-

hibit when feeding and states that once the mouth parts are securely

inserted into the skin the insect is not easily disturbed. Ussova (1961)

states that the mandibles pierce a host's skin, the maxillae with re-

curved teeth anchor the proboscis in the skin, and alternating actions

of the cibarial and pharyngeal pumps suck the blood into the esophagus.





-18-


James and Harwood (1969) indicate simuliids are telmophages or pool

feeders which lacerate blood vessels with the toothed, transversely

operating mandibles and vertically operating maxillae. Davies and

Peterson (1956) record a wide, natural host range for some species such

as S. venustum duck, crow, heron, deer, and human. Others such as

P. hirtipes (now three species) and S. decorum which feed on? mammals in

the field fed on birds when placed in vials on the avian species in

the lab. Yang and Davies (1974) examined the salivary glands of three

black flies and found an anticoagulant factor which keeps the blood

fluid for movement into the gut. These authors also report an agglutin

factor in flies at least twelve hours old.

Adult egg development and egg laying. Cameron (1922) found

development of the ovaries of female black flies followed a successful

blood meal. Davies and Peterson (1956) describe females of several

species with weak teeth or only hairs on the mandibles and maxillae. These

flies do not suck blood and emerge with already ,mature eggs. Davies

and Peterson (1956) indicate eggs develop five to twenty-one days after

emergence with the longer time involving a prolonged blood meal search

under natural conditions. Immature and mature eggs are found together

in the ovaries which suggests at least two ovarian cycles (Cameron,

1922). Females may survive long enough for three batches of eggs but

few probably live this long in the field (Davies and Peterson, 1956).

Cameron (1922) reported that the eggs of Simutium simile (=S. arctioum)

are oviposited on rocks in large cake-like masses embedded in a soft

gelatinous matrix, Davies and Peterson (1956) mention Prosimulium and

Cnephia species which deposit eggs freely while flying down or across

a stream. Stone and Jamnback (1955) indicate S. vittatum lays strings









of eggs in a gelatinouls matrix. Golini and Davies (1975) in Canada

found that female black flies settled at the water line on trailing

cattail leaves and deposited large (16 cm x 2 em) irregular egg masses

one to five layers deep in a gelatinous substance. Davies and Peterson

(1956) found simuliids oviposited an average of 200 to 500 eggs per

female while Golini and Davies (1975) report an average of 417 eggs

for each female.

Adults life cycle, longevity, range, and general activity.

Tarshis (1968) reports a 21 to 25 day period for mixed cultures of five

Simuliaum species to develop into adults after eggs were placed in aquar-

ia. Raybould and Grunewald (1975) found developmental time from egg to

adult ranged from 18 to 50 days for African black flies.

Dalmat (1955) in Guatemala used colored dyes and the release-

recapture technique and found marked female black flies survived as

long as 85 days and traveled as far as 9.7 miles. Bennett (1963a) tagged

females of S. rugglesi by feeding them on ducks injected with a phos-

phorous-32 solution and recovered labelled flies up to 28 days later and

9.6 km? distant following watercourse paths. West-et al. (1968) exposed

black fly larvae and pupae to 32P-treated water and recovered radioactive

flies as far as 33.5 km (20.8 mi) distant. Bennett and Fallis (1971)

found S. eurycdniniculum flew up to five miles from the release point

and report the average life span of the females was at least two to

three weeks. Hocking (1953) summarized published records of flight

ranges which reached a maximum of 145 km for S. reptans columbaczen~se.

Wellington (1974) reported frenzied activity in black flies which

was apparently correlated with barometric pressure changes as a storm

system approached. Cameron (1922) observed swarms of male black flies





-20-


or males and females on warm, cloudy days with rain threatening or

falling gently. Carlsson (1967) reports that rapid changes in tempera-

ture, air pressure, and light seem to increase the activity of all

simuliid species but he also mentions even a light breeze reduced black

fly activity considerably. Hocking (1953) found flight was continuous

in Simulium species down as low as 12.80C (550F).

Concerning nocturnal behavior Dalmat (1955) observed that black

flies in Guatemala move down to the base of plants as the sun sets and

can be captured at night using a lantern and sheet when the insects

emerge from their resting sites near the ground. Wolfe and Peterson

(1960) describe climbing trees and sweep netting twenty-five feet off the

ground to capture black flies at night in Quebec. A4t dawn Wolfe and

Peterson observed simuliids flying down from the canopy. Recently in

India researchers used light traps and found that black flies were active

throughout the night with a peak of flight activity occurring around

midnight (Datta and Dasgupta, 1974).

Lab colonization. Black flies have been collected in nature in all

stages and have been brought to the lab for experiments. Tarshis (1965a)

describes techniques for collecting and shipping viable black fly eggs.

Tarshis (1965b) and Tarshis and Adkins (1971) discuss collecting

large numbers of black fly larvae on artificial cloth substrates and

mention techniques for transporting larvae in aerated containers.

Tarshis (1971) discusses rearing black fly pupae to adults in individual

brass strainer cloth cylinders connected to aquarium air stone units.

Tarshis (1972, 1973) describes field collection, laboratory rearing of

immatures to adults, and successful feedings of females of C. ornitho-

philia on ducklings in the lab.





-21-


Wenk and Raybould (1972) point out that colonies of insects aid

critical studies on their biology, provide adults free from infection

for transmission work and can provide abundant material for investiga-

tions at times when natural populations are limited. Dunbar (1969)

mentions that hybridization experiments supported by colonies of flies

and sufficient knowledge of mating requirements can provide valuable

information concerning the species status of insect forms. Dalmat (1955)

found carbon dioxide stimulated feeding and oviposition and succeeded in

inducing 40% of 65,000 S. ochraceum, S. callidum,; and S. metaieum to

feed and 20% to oviposit but no eggs developed into larvae. Pield et al.

(1967) found that females of S. vitbatum confined in a vial bit man,

quail, and rabbits. By impaling a male on a minute and brushing the

male genitalia against the genitalia of the female coupling could be

achieved. No female of S. vittatum which mated survived to oviposit

and eggs deposited by other, unmated, females in the artificial flows

available failed to hatch. In the laboratory Wenk (1965) successfully

reared, mated, blood fed and achieved oviposition of viable eggs with

Boophthora ery~throcephala. Wenk and Raybould (1972) working with the

Kibwezi form of S. damnrosum likewise reared, mated, blood fed and ob-

tained viable eggs from female black flies in the lab. Mating, which

was also achieved with a member of the S. bovis complex, occurred in

partially lighted emergence cages and was confirmed by the presence of

sperm or spermatophores. Raybould and Grunewald (1975) review the litera-

ture on lab colonization of black flies and mention difficulties confront-

ing researchers: inducing mating, inducing females to feed on blood suf-

ficiently and consistently, erratic viability or hatchibility of eggs

oviposited in the lab, and removing wastes from immature rearing setups.









Although a few black fly species have been induced to complete every

stage of their life cycle in the lab, they are still not considered to

be successfully colonized.



Damage


Jamnback (1973) describes the black fly bite reaction: a hemor-

rhagic spot which develops into an itching wheal; sensitive individuals

may suffer headache, fever, nausea, glandular enlargement, and bites

around the eyes may cause swelling that results in obscured vision.

Loomis et al. (1975) report black flies cause considerable irritation

and tissue damage to horses' ears, head, neck, and belly,producing

wounds and papules. S. archiewn, C. pecwuarn, and S. columbaczense are

described as major pests of livestock (Jamnback, 1973). Fredeen (1974)

mentions black fly outbreaks during 1944-1947 in Saskatchewan which re-

sulted in the deaths of 1100 farm animals including cows, horses, hogs,

and shorn sheep. Fredeen (1975) reports a S. arcticum outbreak in 1972

along the N'orthern Saskatchewan River that killed at least 18 cattle.

Black flies transmit Leucocytozoon parasites to ducks (Fallis and

Bennett, 1966), trypanosomes to ducks (Desser et al., 1975), and

Leucocytozoon parasites to turkeys (Skidmore, 1931; Noblet et al., 1972)

(see also the Leuc-ocytozoon smithi sections in the Literature Review and

Results and Discussion below). Sudia et al. (1975) mention that epi-

demic VEE virus has been isolated from Simuli~um species in Colombia and

although biologic transmission has not been proven, mechanical trans-

mission by contaminated mouth pprts may be possible for at least 72

hours. Travis et al. (1974) report a finding of vesicular stomatitis

virus in black flies in Colombia. Eastern encephalitis virus has been








isolated from a pool of 100 unengorged S. meridionale which indicates

this species may biologically transmit the disease (Anderson et al.,

1961).

Steelman (1976) reports monetary losses in cattle herds due to

worm nodules caused by simuliid transmitted Onchocerca gutterosa amount

to $500,000 each year in Australia. Onchooerca voloulus,the causative

agent of blinding filariasis in mantis vectored by black flies and has

been reported from Africa, Yemen, Mexico, Guatemala, Venezuela, Colombia,

and Surinamf; indigenous cases may also occur in Brazil (World Health

Organization, 1971; Travis et al., 1974; Raybould and Grunewald, 1975).

i~crofilaria of 0. voloulus previously were observed in the eyes and

skin, however, Anderson et al. (1975a) found microfilaria in the urine,

blood, and sputum of persons treated with diethylcarbamazine. Diurnal

periodicity of 0. volvulus microfilaria in the skin has been shown to

correspond with the peak feeding periods of important black fly vectors

in Guatemala and Africa (Anderson et al., 1975b). Duke et al. (1975)

found the transmission potentials of 0. volvulus along breeding rivers

for S. d'awnosum in Africa so high that no communities could survive there.



Control


Physical control. Impoundage of rivers and the creation of reser-

voirs together with planned periodic cutoffs of discharge water has

discouraged or eliminated black fly breeding along many stretches of

rivers (Snow et al., 1958). Removing debris such as planks, tree limbs,

trailing vegetation and other potential substrates for black fly larvae

can help control s~imuliid population levels in streams (Jamnback, 1973).









Chemical control. Carestia et al. (1974) reported that serial ap-

plications of malathion, DibromR or Dibrom plus a repellent for

adult black fly control were unsatisfactory. High winds, dropping

temperatures and decreasing daylight interfered with adult fly activity

and chemical effectiveness. Rapid reinfestation of adults from areas

just outside the treated zone is another reason why most control efforts

have been aimed at immature rather than adult black flies.

Fairchild and Barreda (1945) observed the effectiveness of DDT as

a black fly larvicide. Davis et al. (1957) discussed early attempts at

evaluating the larvicidal properties of parathion dripped into streams,

dieldrin applied by air, and DDT delivered by various ground techniques

as well as by airplane. Evidence of the persistence of DDT in non-target

organisms and decreased susceptibility of black fly larvae from streams

with a history of DDT treatments stimulated a switch to less persistent

materials (Fredeen et al., 1971; Jamnback and West, 1970). Travis et al.

(1970) in field trough tests found Dibram was outstanding in producing

mortality of S. pictipes larvae at .5 ppm and 1 ppm concentrations and

stated Dibrom, GardonaR and CiodrinR deserved more practical stream

tests.' Jamaback and Frempong-Boadu (1966) found chemical formulations

are most effective which give uniform distribution of the insecticide

in the water and have a specific gravity of slightly less than 1.0 to

keep the toxicant near the surface where most of the black fly larvae

are located. AbateR (20%) in Panasol plus .5% Triton X-161 (.998

specific gravity) applied by aircraft eliminated larvae up to .8 km

(.5 mi) downstream from the treatment point. Pelsue et al. (1970) chose

Abate as a black fly larvicide due to its low mammalian, avian, and fish

toxicity and achieved 100% control of larvae in three locations









with .5 ppm delivered for 60 minutes. Detachment of larvae occurred

within 24 hours and reinfestation was noted in 15 to 60 days. Kissam

et al. (1975) reported 2% Abate Celatom granules delivered at 91 g AI/

.4 ha monthly reduced larval populations t~o zero and the populations

only slowly built back up during the two weeks following each applica-

tion.

Fredeen (1974) mentions that methoxychlor, another black fly larvi-

cide, is minimally toxic to vertebrates and methoxychllor and its

metabolites are not concentrated in fish or other aquatic species. In

Canada in 1969 a 15 minute injection of .2 ppm methoxychlor emulsifiable

concentrate caused the disappearance of 96% of the S. arcticum larvae

32 km downstream. Fredeen (1975) reported that a 7.5 minute injection

of methoxychlor at .6 ppm killed 100% of the older black fly instars 40

and 80 km below the treatment area. Younger black fly instars which are

more susceptible were depleted by 96% at a site 161 km downstream from

the injection. Fredeen et al. (1975) indicate that methoxychlor adsorbs

to suspended solids in the water and may act selectively against the

filter-feeding black fly larvae. Following a methoxychlor application

Wallace et al. (1973) noted increased drift but no eradication of non-

target organisms and Fredeen (1975) found that non-target organisms

repopulated more densely than before the treatment.

Investigations of the effect of insect developmental inhibitors on

black flies indicate that significant reduction (75-100%) of adult

emergence can be attained in the lab with at least three black fly

species with AlcosidR at concentrations between .001 and 1 ppm (McKague

and Wood, 1974; Dove and McKague, 1975). Frormmer et al. (1975) reported

that in field evaluation tests in New York DEET-treated light mesh





-26-


jackets were effective in reducing landings per five minutes from 404

on controls to 1.2 on personnel with treated jackets.

Biological control. Cameron (1922) found that a fish of no food

value called the common sucker fed on S. simile (=S. arcticum) larvae

and pupae. Peterson and Davies (1960) review the predators of black

flies which include adult Empididae and Dolichopodidae, Tendipedidae

larvae, Trichoptera, Formicidae, Odonata -adults and naiads, and spiders.

Sommerman (1962) reported empidid larvae fed on black fly larvae.

Snoddy (1968) found the solitary wasp Oxrybelus emarginatum to be a

predator of adult black flies. Peterson and Davies (1960) and Chutter

(1972) report cannibalism among simuliid larvae. Carlsson (1967) in-

dicates predators are unlikely to serve well as man-manipulated biologi-

cal control agents though the tricopteran genera Hiydropsyche and

R~hyacophila might give good results.

One of the earliest accounts of parasitism of black flies is that

.of Strickland (1911) who reported a mermithid nematode that retarded

the development of the larval histoblasts and a sporozoan (microspori-

dian) which caused distorted and swollen larvae. Jamnbackc (1973) states

that eighteen microsporidia have been described from black flies.

Spores are ingested by the larvae, sporoplasm invades suitable host

cells which are often in the fat bodies, multiplication and the pro-

duction of many spores occurs, and the host larvae commonly die. Black

fly pupae and adults have been found infected and transovarian trans-

mission is common. Jamnback (1973) also mentions a fungus Coeiomycidiwn

simulii which occurs in black fly larvae and is usually fatal.

Davies (1958) reports microsporidians, mites, and mermithid nema-

todes as parasites of Canadian black flies and mentions that mermithids









are found in larvae, pupae and both sexes of the adults. From 15 to

60% of the females in emergence collections were found to be infected

with mermithids and females in oviposition swarms were also found with

mermithids. The infected females which attempt to oviposit may serve to

disperse the mermithids and introduce them to additional stream environ-

ments. Welch (1964) indicated that the mermithid life cycle began with

the consumption by black fly larvae of the mermithid, usually in the

infective first juvenile stage. Molloy and Jamnback (1975) observed

direct penetration of the black fly larval cuticle by preparasitic

juveniles of Neomesomermis fluiminais, a mermithid which infects at

least fourteen species of black flies in three genera. Exit of the

parasite from the host is almost immediately fatal to the black fly.

Anderson and DeFoliart (1962) report 49-93% parasitism by Isomermis and

Gastromermis in one black fly species in Wisconsin. As a step toward

the mass rearing of these biological control agents Bailey et al. (1974)

discuss techniques for the mass collection of mermithid postparasites

from field collected black fly larvae.

Batson et al. (1976) reportedatiridescent virus from black fly

larvae in Wales but could associate no major fluctuations in the black

fly population with the presence of the infected larvae.



Florida Ecological Habitats


Florida is a peninsula that lies between 80 and 88 degrees west

longitude and in the same latitude belt as the Sahara, Arabian, and other

large deserts. Exclusive of the Keys, the State extends roughly 644 km

(400 mi) north and south along the peninsula and about the same distance

east to west along the north coast of the Gulf of Mexico. Florida covers





-28-


151,710 sq km (58,560 sq mi)'. The highest point in the State is 105 m

(345 ft) above sea level, at Lakewood in Walton County. From Orlando

south to Sebring another high section occurs and Iron Mountain near

Lake Wales is located about 91.5 m (300 ft) above sea level (Morris,

1973). The earliest geological horizon is the Ocala limestone formed

during the Eocene, 58 million years ago. Modern Florida made its first

appearance during the Oligocene, 36 million years ago, as a small island

about where the counties of Suwanee, Columbia, and Alachua are now

located and then rose as a peninsula during the Miocene (Byers, 1930).

There are three schemes which have been used to break Florida up

into sections according to: 1) vegetation; 2) geology; and 3) edaphic

factors. These schemes are presented to help illustrate the ecological

diversity in Florida, which an the one hand provides a range of habitats

for simulfid species and on the other hand restricts black fly breeding

to certain more suitable regions of the state. Byers (1930) divided

Florida into seven biotic areas based on dominant types of vegetation:

a tropical hammock strip from St. Lucie County through Dade County along

the east coast extending a few miles inland with cabbage palms, mahogany,

ironwood, papaya, epiphytic bromeliads, plus other plants; a grass swamp

area south and east of Lake Okeechabee; magnolia and temperate hammocks,

primarily along the east coast from Flagler County to Indian River

County and on the Gulf Coast from Wakulla through Hernando Counties -

these occur on well drained but moisture holding soils with magnolia,

holly, and red bay as dominant vegetation on the east coast and beech,

elm, sweetgum, hickory, and oaks dominant in the west coast hammocks;

a section of southeastern deciduous forest extending down into Leon,

Liberty and Jefferson Counties; tree swamps in Collier, Columbia, Baker









and Nassau Counties which include Big Cypress Swamp and other cypress

and also black gum swamps; pine flatwoods along the Gulf west of

Wakulla County, on the west coast of the peninsula from Pasco to

Collier Counties, and in the Clay County and Putnam County region -

these flatwoods occur on level, poorly drained land underlain by hard

pan which results in an acid soil upon which grow palmetto, grass and

pines; and, lastly, the southeast coniferous forest with long leaf,

yellow, and slash pines and saw palmetto on well drained uplands.

Cooke(1939) divided Florida into five physiographic areas: central

highlands, Tallahassee hills, western highlands, Marianna lowlands and

coastal lowlands. The central highlands, in the center of the peninsula

from Baker County and Columbia County south to Lake Okeechobee, contain

in their southern section thousands of lakes which in the north section

are accompanied by numerous streams. The Tallahassee hills and western

highlands are composed mainly of red clay with a relatively good number

of streams. The coastal lowlands usually lie less than 30 m (100 ft)

above sea level and include swampy areas such as the Florida Everglades

and marshy areas along the east and west coasts of the state. The

Marianna lowlands in Wlalton, Holmes, Washington and Jackson Counties

contain few permanent flows and consist mainly of swamps, flatwoods,

and sandy hills bearing pines.

Davis (1967) and Smith et al. (1967) present six major land resource

areas for Florida: a southern coastal plain; gulf coast flatwoods;

central Florida ridge; Atlantic coast flatwoods; southern Florida flat-

woods; and Everglades and associated areas. The southern coastal plain,

occupying the northern half of the state from Escambia County to Madison

County,is covered by forests of mixed hardwoods and pines on the lower





-30-


areas and forests of long leaf pines and oaks on upland clay or well

drained upland sand. The gulf coast flatwoods--a strip along the gulf

side of the panhandle from Escambia County through Citrus County, the

Atlantic coast flatwoods primarily in Columbia, Baker, Nassau andBrad-

ford Counties in the north, and the south Florida flatwoods occupying

most of the southern section of the peninsula outside of the Everglades

consist of pine flatwoods and swamp forests (pines, palmetto, herbs,

bay tree, laurel, gum, cypress) on poorly to very poorly drained and

marshy soils. The central Florida ridge is located on well drained

soil in the middle of the peninsula and includes forests of long leaf

pine and xerophytic turkey oak (now mainly planted in citrus) and hard-

wood forests of mixed evergreen and deciduous hardwoods on rich upland

soils. The Everglades are primarily sawgrass, M~ariscus jamaioensis, on

peat and muck soils.

Average January temperatures range from 10-130C (50-550F) in the

panhandle to 18-210C (65-700F) in the Everglades area. Average July

temperatures fall between 27-290C (80-840F) throughout the state. An-

nual rainfall averages 134.62 cm (53 in) (Raisz, 1964). June through

September or October is considered the rainy season in Florida (Byers,

1930) however Berner (1950) indicates that in the northwestern area

rainfall is more evenly distributed throughout the year. There are

seventeen first magnitude springs in Florida (Morris, 1973). Despite

the heavy precipitation there are relatively few surface flows since

much of the water moves in underground channels in the underlying lime-

stone (Raisz, 1964). The vast majority of true streams and rivers are

found north of L~ake Okeechobee. Morris (1973) indicates there are 1,711

streams, rivers, and creeks in Florida with a total length of 16,989 km

(10,550 mi).








Berner (1950) states that in Florida there are relatively few

intermittent streams. He divides permanently flowing streams in the

State into: sand-bottom creeks with loose shifting sand and little

vegetation; sand-bottom creeks with fairly loose sand and considerable

vegetation; silt-bottomed creeks with little vegetation; and silt-

bottomed creeks with considerable vegetation. Stagnant rivers occur in

south Florida where they serve primarily as drainage canals for the

Everglades and though there is considerable vegetation along the shores

of the rivers their fifteen to twenty-foot depths restrict plant growth

toward the middle. Larger calcareous streams contain water which rises

from springs, are basic in pH reaction, and in shallower sections are

lined on the bottom with Vazllisneria (eel grass), Sagittaria (arrowhead),

algae and other plants. Erving (1971) notes that the temperature for

Florida springs is about 220C (720F) year round. While noting springs

are alkaline Berner (1950) also indicates that marsh and swamp water can

be very acidic reaching a pH of 3.6 or below. He further adds slow

flowing rivers to his classification of Florida's streams and rivers

and included here? are the Suwanee, Apalachicola and other rivers which

have extensive drainage areas. Wood and Fernald (1974) list twenty-

eight major drainage basins and streams in Florida. The two largest,

the Apalachicola River Basin and the Suwanee River Basin drain 50,777

sq km (19,600 sq mi) and 25,984 sq km (10,030 sq mi), respectively,

some of which lies outside of Florida. Wood and Fernald (1974) indicate

that peak annual flows occur in eastern rivers in September and October

with buildups since June while more western rivers (Chipola, Apalachi-

cola) reach maximum flow during March and April. In northwesternl

Florida a continental weather pattern predominates whereas a more









tropical weather routine predominates in central and south Florida.

Beck (1965) recognized five stream types in Florida: sand-bottomed,

calcareous, swamp and bog, larger rivers, and canals. He points out

that due to velocity, substrate, and spring discharges many of these

types of flows can be recognized at different points along a single

watercourse and cites the Suwanee River which is successively a swamp

and bog stream, a sand-bottomed stream, a calcareous stream and a larger

river of mixed origin. Beck (1965) classifies swift flow in Florida's

streams as that velocity suitable for Plecopterawhile simuliid popula-

tions indicate conditions of moderate velocity. Below these two levels

the speed is considered low. A general progression is recognized with

the swifter portions of upper rivers marked by eroding limestone and

thick growths of moss, midpoints along rivers being sand-bottomed and

lower, slower portions of rivers containing deposits of finer materials

on the bottom. Beck (1965) lists the sand-bottomed stream as the most

common type of stream in Florida. The fauna consists of Hydropsychid

and Philopotamid caddistlies, simuliid larvae, Ple-coptera, Stenonema

mayflies, orthocladine chironomids, and Corydatis. In these sand-

bottomed streams the pH reaction is usually between 5.7 and 7.4 and

they display moderate to high color and moderate to swift velocity. The

bottoms consist of sand, leaf deposits, and limestone outeroppings, and

plant growth may be dense. Beck mentions that swamp and bog streams are

highly acidic (pH reaction 3.8-6.5), sluggish flows found in the coastal

lowlands and central highlands with origins in swamps and marshes.

Calcareous streams are cool, clear flows of spring origin with dense

growths of vegetation. These flows are alkaline (7.0-8.2 in pH), bear

large mollusc populations, are generally of low color, of variable










velocity and have sand, clay or limestone bottoms. The flows in the

larger river category usually carry considerable silt and are turbid,

have high and steep banks and bottoms of coarse sand and limestone.

They demonstrate a pH reaction of 6.5-7.4 and have few shallow places

and few aquatic plants. Beck (1965) mentions from about the area of

the St. Lucie Canal to south of Homestead no natural streams of any

consequence remain along the east coast of Florida and, instead, canals

are prevalent although in the past the Miami River had a swift flow and

rapids (Fairchild, 1976 personal communication).

The above discussion provides an introduction to the climatic,

edaphic, biotic and hydrologic conditions which are found in the State

of Florida.



Leucooptozoon smith


During the 1880's Danilewsky observed parasites which were later

described and designated as species of Leucocytozoon and Haemoproteus

(Bennett et al., 1965). Hsu et al. (1973) list 67 species of

Leucocytozoon, one of which is reported from Meteagris gallopavo, the

domestic and wild turkey. Theobald Smith in 1895 first reported a

protozoan-parasite in the blood of turkeys in Massachusetts and Rhode

Island (Smith, 1895). Similar gametocytes were observed in the blood

of turkeys in France and named Haemamoeba smith (Laveran and Lucet,

1905). The following taxonomic scheme which shows the position of the

turkey parasite now known as Leucocytozoon smith was modified from

Aikawa and Sterling (1974) by crediting Sambon (1908) not Danilewsky

(according to Bennett et al., 1975) with the genus Leucocytozoon:








Phylum Protozoa Goldfuss, 1818, emend. von Siebold, 1845
Subphylum Sporozoa Leuckart, 1879
Class Telosporea, Schaudinn, 1900
Subclass Coccidia Leuckart, 1879
Order Eucoccida Leger and Duboscq, 1910
Suborder Haemosporina Danilewsky, 188i5
Family Leucocytozooidae Fallis and Bennett, 1961
Genus L~eucocytozoon Sambon, 1908
Species smith (Laveran and Lucet, 1905).


The following authors observed L. smith in turkeys in their respec-

tive states or countries: Atchley (1951), Bierer (1954), Jones and

Ricbey (1956) and W~ehr (1962) in South Carolina; Savage and Isa (1945)

and Bennett et al. (1965) in Canada; Volkmar (1929) in Minnesota and

North Dakota; Hinshaw and McNeil (1943) in California; Johnson (1945) in

Michigan; Kozicky (1948) in Pennsylvania; Skidmore (1931) in Nebraska;

Stoddard et al. (1952) in Georgia; and Byrd (1959) in Virginia. Volkmar

(1929) also lists L. amithi from Germany and the Crimean Peninsula and

Cook (1971) lists Asia. Travis et al. (1939) reported Leucocytoaoon

smith from wild and domestic turkeys in Florida as well as Georgia,

Missouri, Alabama and South Carolina. Simpson et al. (1956) and Forrester

et al. (1974) also reported Leu~cocytozoon smithi from Florida turkeys.

Solis (1973) exposed turkeys, ringnecked pheasants, chickens, bob'white

quail, pekin ducks and chukar partridges in an area where L~. snrithi

occurred and found that only turkeys were susceptible to L. smithi.

Fallis et al. (1974) and Aikawa and Sterling (1974) present in-

formation an the life cycle and structure of the intracellular parasite,

L. smith. Leucacotozoon belongs to the same suborder, Haemosporina,

as Plasmodiun and Hiaemoproteus and undergoes a similar life cycle.

Unllike PLasmodium, however, there is no eryrthrocytic schizogony and in

addition to the red blood cells being invaded by merozoites as in

Haemoproteus (and Plrsmrodiuw)lin L~aucocytozoon white blood cells are









also used as sites for gametogony (Huff, 1942; Fallis et al., 1974).

Gametocytes, in the case of L. smith, grow and split the host nucleus

in two (Sambon, 1908; Volkmar, 1929). Plasmodium species are tran~s-

mitted by mosquitoes and Haemnoproteus (Parahaemoproteus) species by the

Hippoboscidae and Ceratopogonidae (Fallis and Bennett, 1961a and b;

Bennett et al., 1974). Leucocytozoon, is known at present to be trans-

mitted only by black flies. A parasite of chickens which is vectored by

Culicoides was designated Akiba caulleryi by Bennett et al. (1965) but

Akiba has recently been considered as a subgenus of Leucocytozoon (Hsu

et al., 1973; Fallis et al., 1974). Gametocytes are ingested with the

blood of an infected turkey by a feeding black fly. Roller and Desser

(1973) observed diurnal periodicity with L. simondi where peak gameto-

cytemia corresponded with peak activity and biting periods of the vector.

It has been suggested that there is periodicity of the gametocytes of

L. smith in the peripheral blood and correlation with the periodicity

and feeding peaks of the vectors (Moore et al., 1974). Early in the

infection each gametocyte appears as a round body in the parasitized

cells; later the gametocytes deform each blood cell into the charac-

teristic spindle shape (Huff, 1942). Desser et al. (1970) found with

L. sim~ondi elongated cells resulted when merozoites developed in leuco-

cytes and round cells resulted from invasion of red blood cells. Macro-

gametacytes stain more darkly with Glemsa stain than microgametocytes.

Within minutes of ingestion the microgametocytes and macrogametocytes

escape from their host blood cells. The microgametocytes exflagellate

and syngamy occurs with the macrogametes. The zygotes formed grow into

elongate ookinetes within twelve hours after the initial blood ingestion.

Sections of flies have revealed ookinetes in the process of penetrating








the midgut of the fly (Fallis et al., 1974). Fallis and Bennett (1961a)

working with L. simonzdi detected only sluggish movement of ookinetes

and expressed doubt that this motion could facilitate penetration of

the gut wall of a black fly. Howard (1962) working with Plasmodium

gallinaceum1 observed no mobility or active penetration by zygotes.

Howard suggests a largely passive incorporation into the gut wall to

within 5 microns of the external basement membrane as the blood meal is

digested and the distended gut returns from a squamous cell configura-

tion to its original columnar cell morphology. Desser (1970) described

a pore, strengthening struts, microtubules and elongate convoluted

micronemes in the apical cap of the anterior end of L. simondi cokinetes

and suggested these structures might aid penetration through the peri-

trophic membrane and into the fly midgut epithelium. In as little as

48 hours the ookinetes develop into oocysts containing 50-100 sporozoites

just beneath the outer membrane of the gut. Sporozoites escape from the

oocysts and penetrate and collect in the salivary glands. Desser (1970)

describes structures on the anterior end of sporozoites of L. simondi

which may contain proteolytic enzymes and aid in entering the salivary

glands and, later, host liver cells. The sporozoites pass out the

proboscis of the black fly with the salivary fluids when the fly bites

another host. The sporozoites of L. sm~ithi invade the liver parenchymal

cells of the turkey and grow into hepatic schizonts. Huff (1942) re-

ports with other species of Leucocytozoon megaloschizonts in the heart,

spleen and other host tissues. Such schizonts are not normally observed

with L. smithi; however Siccardi et al. (1974) reported finding magalo-

schizonts in the kidneys of infected turkeys fed coccidiostat medication.

The schizonts by asexual multiplication (schizogony) produce merozoites





-37-


which enter red and white blood cells. Peters (1971) suggests that in

the related genus Plasmodium merozoite penetration may involve proteolytic

enzymes and be active or may be more passive with the merozoites being

engulfed or invaginated into the blood cells. The merozoites grow into

gametocytes in the blood cells and the cycle is completed. The prepatent

period or time elapsed between entrance of the sporozoites and appear-

ance of gametocytes in the peripheral blood is 10 to 16 days.

Symptoms associated with leucocytozoonosis include: loss of

appetite, emaciation, wheezing breathing, congested heart and lungs,

drooling, drooping wings, sitting on the hocks, enlarged liver and

spleen, anemia, impaired imrmunalogical responsiveness, and death (Wehr,

1962; Salsbury, 1971; Siccardi et al., 1974). Turkeys under 12 weeks

of age are often severely affected but mortality in older birds has

also been reported (Simpson et al., 1956). Many birds with high para-

sitemias appear outwardly normal. Birds that do not die may continue

to carry the parasite for as long as 14 years (Skidmore, 1931). Borg

(1953), Simpson et al. (1956) and others suggest that the pathogenicity

of L. smith has not been proven conclusively and that L. smith may be

an additive debilitating factor which when combined with other factors

such as blackhead, fowl cholera or leukosis results in bird mortality.

Stoddard at al. (1952), Skidmore (1931), Savage and Isa (1945) and

others however have observed flock mortality up to 75% where bacterial

cultures and other tests revealed L. smith as the sole disease organ-

ism present. Jones and Richey (1956) indicated annual death losses due

to Leucocytozoon in one county in South Carolina averaged 5%. In

1973,132 million turkeys were raised in the U.S. and farmers grossed

$934 million (Poultry Digest, April 1974; Agricultural Situation,








November 1974). In the absence of severe threats by disease this indus-

try will continue to grow as the public's demand for poultry and its

products like turkey rolls and t.v. dinners as an alternative to beef

increases.

A number of black flies have been incriminated as possible vectors

of L. smith. Skidmore (1931) incriminated S. occidentale (=meridionale)

as a vector of L. smith in Nebraska by grinding up a number of f-lies

that had fed on infected turkeys, injecting them in saline into clean

birds, and 12 days later observing gametocytes of L. smith in the tur-

keys' blood streams. Johnson et al. (1938) in Virginia exposed uninfected

turkeys to the bites of S. nigroparvum (=jenningsi) and obtained

L. amithi transmission. Underhill (1944) further substantiated this

work but Byrd (1959) was unable to infect clean turkeys by macerating

and injecting S. jenningsi that had fed on diseased birds. Wehr (1953)

by injection and Jones and Richey (1956) by fly-bite incriminated

S. skossonae as a vector of L. smith. Byrd (1959) succeeded in

infecting turkeys with L. smith by grinding and injecting females of

Prosimulium hirtipes that had fed on infected birds. I have seen no

further follow up of this work and Fallis et al.(1974) do not list any

member of the former P. hirti~pes complex as a vector of L. s ithi.

Noblet et al. (1972) ground up in saline and injected infected females

of S. congareoenruAm into turkeys and incriminated this species as a

suitable or possible vector for L. smith. Savage and Isa (1945) and

Anthony and Richey (1958) have reported leucocytozoonosis in the sup-

posed absence of simuliids although other biting flies such as CulcZcoidesJ,

Stomorys, Di~achlorus and mosquitoes were present.

Control of leucocytozoonosis can be achieved by locating :urkey

flocks at least 12.8 km (8 mi) from known breeding locations of potential









vectors and in areas free from possible wild turkey disease reservoirs.

Control of black flies by large scale aerial treatment of streams with

larvicides has been shown to markedly decrease the level of

parasitemias of L. emithi in sentinel turkeys (Kissam et al., 1975).

Fine mesh wire screen enclosures might be a valuable preventive tech-

nique with smaller flocks. Clopidol, a coccidiostat, at .025 and

.0125% in feed is effective in reducing the number of L. smithi para-

sites in turkeys as indicated by blood smears (Siccardi et al., 1974).

Investigators of leucocytozoonosis in turkeys in Florida have

studied the presence, consequence, and distribution of L. smithi in

the State but have not proven specific vectors. Travis (1939) reported

that nearly 100% of the wild turkeys in low areas of Georgia and Florida

were infected with L. smithi and suggested that an aquatic breeding

insect might be involved. Simpson et al. (1956) investigating out-

breaks of L. smith in turkey flocks near Palatka, Florida, reported

finding S. slossonae and a species #158 (=S. jonesi Stone and Snoddy,

1969) breeding near the turkey flocks and suggested that S. slossonae

might be an important vector. Davis et al. (1957) reported S. slossonae

as prevalent in Florida. Forrester et al. (1974) found a 72-75% in-

cidence of L. smith in 484 turkeys more than one month old sampled

from 10 localities in Florida. These authors also noted that a drop in

the infection rate of L. smithzi observed in wild turkeys in the Sikes

Fisheating Creek Wildlife Management Area and other areas of the State

in 1971 corresponded with low rainfall and low stream conditions and

theorized that reduced numbers of potential black fly vectors may have

resulted in the reduced prevalence of Leucocytozoon in turkeys.














CHAPTER III
MATERIALS AND METHODS



Black Fly Survey


Primarily, efforts in this survey concentrated on collecting larvae

and pupae from Florida's streams and rivers. In Alachua County ten

streams were selected and in nearby counties an additional twenty flows

were chosen which were fairly permanent and represented a diversity of

ecological conditions. An attempt was made to secure samples from each

of these streams at least once a month. Approximately the same location

was revisited each time to obtain some consistency in stream and sub-

strate conditions in order to judge seasonal replacement phenomena and

normal population changes. Other sites in south, southwest and west

Florida were chosen, generally along highways or country roads for ac-

cessibility, which were visited at least quarterly to obtain records at

all seasons of the year.

At each location specimens were removed with forceps or placed

still attached to small portions of grass, twigs and so on into four-dram

lip vials with neoprene stoppers. Each vial contained 80% ethyl alcohol

and a code number penciled on a small slip of paper. The immatures,

collected in 30 to 45 minutes were deemed sufficient to indicate numbers

and species present. Dark pupae, indicating an advanced state of develop-

ment, were placed in four-dram screw cap vials on a strip of paper toweling

moistened in the stream and were held alive in order to allow the adults

to emerge. Images could be reared very successfully in this manner from





-41-


older and frequently, even younger, more pale pupae. The forceps used

were secured to my wrist by a string to prevent their loss when my

balance was upset by the current, when I was wading and saw an occasional

water moccasin, and at other critical times. A 1.53 m long rake and a

modified, extendible, golf ball retriever were used to obtain substrate

samples and specimens where it was impossible to wade.

At each collection location stream dimensions, velocity, pH, tem-

perature, color, and substrate were recorded an special data cards along

with indications of the attachment substrates of the immatures and an

estimate of the population size. Temperatures were obtained with a

metal-cased pocket thermometer, range -lo to 490C (300 to 1200F). Initial

attempts were made to determine the pH of the streams using two portable,

electronic pH meters. After obtaining erratic, inconsistent results in

the field with the meters, pH papers (pHydrion papers Micro Essential

Labs, New York) were tried with more success. These pH papers come in

ranges such as 3.5-5.5 and 6.0-8.0 between I and 12 pH units with illus-

trated color differences at .5 pH unit intervals and are rated accurate

to .25 pH unit in buffered solutions. The papers were checked regularly

for accuracy with pH 4.0, 6.5, 7.0 and 10.0 buffered solutions. Very

acidic readings were confirmed by bringing chilled water samples into

the lab and checking them on a Beckman pH meter. Experiments with

electronic, propelleredGurley flow meters proved them too cumbersome to

set up and difficult to use in the streams which were often shallow and

clogged with submerged vegetation. The floating cork technique suggested

by Dalmat (1955) and others was used. A weighted cork 3.8 cm in diameter

which floated low in the water and was connected to a 3.05 m (10 ft)

long string was timed with a stop watch to determine the stream velocity









at different points where larvae and pupae were located. Most immature

black flies were encountered on trailing vegetation at or near the sur-

face and thus the floating cork velocity figures should reasonably

reflect immature habitat conditions. A ruler one meter long with

centimeter markings was used to determine stream depths and widths. The

widths of larger flows were estimated by pacing across them, when possi-

ble, or by pacing across a nearby bridge. Other omnipresent items on

collecting trips included an insect net an-d a snake bite kit.

Alcohol specimens and pupal vials from each site were placed with

the data card into cloth bags and transported to the lab where identi-

fications were made with the aid of a Bausch and Lomb stereamicroscope.

Larvae were usually identifiable without dissection (except unraveling

of the respiratory histablast); however representatives of each species

were dissected and mandibles, respiratory histoblasts, cephalic apotome,

gular notch, antennae and fans, and anal sclerite and hooks were pre-

pared in cellosolve and mounted in balsam under individual small cover

slips. Adults which emerged were allowed to harden for a few hours and

were then killed and placed in alcohol, prepared and mounted in balsam

on slides, or pinned on minutin nadeln inserted in white polyporous

pieces on #3 insect pins.

In addition to immatures, some adult collecting was undertaken

using a variety of traps. Black fly adults have been obtained in Florida

by other individuals using light traps and Malaise (flight) traps. In

this research three types of adult fly traps were used: the Manito'oa

trap (Thorsteinson et al., 1965), the blackout box trap (Anderson and

Dicke, 1960), and a modified ramp-type trap. The latter two types of

traps will be discussed with the Leucocytozoon. work. A modified Manitoba









trap (Fig. 1) was constructed using 3 bamboo poles, each 2.1 m (7 ft)

long, with a hole drilled vertically into the uppermost node of each

pole into which a leg of a metal tripod laboratory ring stand was in-

serted. A triangular serex or organza canopy 1.2 m (4 ft) tall was

positioned inside the leg frame and anchored at the three lower corners

by strings tied through a hole in each pole. The apex of the tapering

canopy with a 5.1 cm (2 in) wide elastic opening was drawn up a short

distance through the ring opening and stretched around the lower lip of

a 17.8 cm (7 in) tall lantern jar, the collecting vessel, which rested

on the ring. Th-e jar was topped with a double layer of white serex or

organza held in place with rubber bands. The jar contained an inverted

plastic funnel affixed to the jar with hot plastic glue. On strings

attached around the base of the jar and hanging into the canopy was

suspended a 1.2 m (4 ft) long cord with a black, cylindrical (20 cm

wide, 22 cm tall) metal can or a black cardboard triangle, 63 cm on

each side, on the end. A 23 cm wide black plastic "skirt" was sometimes

fitted over the top of the canopy and positioned at the lower margin of

the canopy to accentuate the "window" effect. A 31 cm long cloth bag

which contained 1.36-1.8 kg of dry ice was also hung inside the canopy.

The dry ice and the black target attracted Simuliidae and other Diptera

which flew or walked up the canopy into the collecting jar. Adults

could be aspirated from the jar by removing the outer cloth layer and

inserting an aspirator tube through a small slit in the lower layer of

material. Specimens could also be taken to the lab in the glass jar and

immobilized by cold in a refrigerator, though condensation inside the

jar often wet the specimens. Manitoba. trapping was accomplished using

one or two traps at a time in six counties at twelve locations for one





-44-


C'? -

'~.., $~
~~~~
r~b-i~~a a


Figure 1. A modified Manitoba trap with a black plastic skirt.









to eight hour periods during the months of January through October. The

air temperature, wind speed and relative humidity were measured during

the collecting periods. Trapped adults were pinned or mounted on

slides as with reared adults. Identification of irmmatures and adults

was possible using taxonomic keys in Stone and Snoddy (1969), Stone

(1964) and smaller publications on individual species.



Leucoc~ytozoon amithi Transmission


In the investigation of the transmission of L. smith to turkeys

in Florida, modifications of Koch's postulates were used as guidelines.

The first aim was to establish an association in nature between the

host, the disease, and the vector. It was necessary to have a sus-

pected vector, preferably a clean, reared one, feed on an infected host

in the lab and to observe the stages of the parasite, especially the

infective stage, the sporozoite, in the fly. A clean host was then to

'be infected by a vector which had become infected in the lab with the

Leucocytozoon parasite being recovered in the gametocyte stage from the

formerly uninfected turkey.

In order to confirm the reported presence of L. smithi in certain

areas such as the Lochloosa Creek and Fisheating Creek Wildlife Manage-

ment areas (L. Williams, 1974 personal communication; Forrester et al.,

1974), I set out sentinel turkeys. The same was done to establish the

presence of the disease in other areas, and to obtain infected hosts to

serve as donors for transmission studies. These turkeys, like all clean

poults used in the transmission studies, were either hatched from fertile

eggs in a fly-proof room or obtained as disease-free one-day old poults

and held in a fly-proof room until use. Difficulties in obtaining clean










poults for experiments early in 1975 when winter and early spring species,

suspected to be L. smithi vectors, were flying were overcome for work

in early 1976. Usually three birds were placed out in the field near

a black fly stream in a cage for about a week and provided with food

and water (Fig. 2). There were three primary sentinel sites, all in

Alachua Co., and additional birds were also set out for short periods

at Fisheating Creek, Glades Co. (see Table 5). Sentinels were also

placed out on a few occasions in a ramp trap described below. In addi-

tion I worked with Dr. D.F. Forrester at Fisheating Creek during 1974

when he exposed many young turkeys for two week periods in cages in

cypress trees and on the ground and in 1975 when cages and ramp traps

were used for turkey exposures.

To determine the species of black flies generally present at the

sentinel sites, immature stages were collected and adults on the wing

were sampled with Manitoba traps as described in the previous section.

To identify black flies specifically attracted to turkeys I used blackout

box traps (Fig. 3), ramp traps (Fig. 4) and exposed turkeys (Fig. 5).

Anderson and DeFoliart (1961) used the technique of blackout box trapping

in Wisconsin. In Florida the blackout box traps were used at two

locations in 1974 and an additional seven sites during 1975 (see Table 7).

A turkey was exposed in a cage on the ground or on a platform in a tree

for about 15 minutes. A large cardboard box with two plastic collecting

cups on the top which appeared as bright areas inside the otherwise dark

box was then placed over the cage and bird. Black flies completing their

blood meals or unfed but on the turkey exhibited a positive phototactic

response, flew to the collecting cups and were captured. The flies were

removed to holding cartons, the turkey was re-exposed, and the process

was repeated.





-47-


;.j13 '' ~k9- );~x~i~


Figure 2. Sentinel turkeys in an exposure cage.


Figure 3. A blackout box trap in the field.





-48-


Figure 4. One view of a ramp trap.


Figure 5. An exposed turkey in the field.


uirPLP)iC' n






t I (1~

1]L~





-49-


An omnidirectional ramp trap was used with the hopes that black

flies would be captured without the constant presence of the collector.

The trap was used at three locations from April to July 1975 (see Table

6). The ramp trap was a wooden-framed cube approximately .92 m (3 ft)

in each demension with 4 slanted lower level ingress panels of aluminum

framing and organza material and 4 upper, vertical organza-paneled

windows. A cage with host animals was placed and suspended inside the

trap through a hinged access opening cut into the solid plywood top.

The attracted insects would fly or bounce up the slanted ingress open-

ings, enter the trap to feed on the hosts, theoretically be unable to

escape and later be aspirated from the inside of the bright vertical

cloth panels or windows.

By capturing flies off normally one or two exposed turkeys it was

possible to tell which species approached the birds and, more signifi-

cantly, to observe which species fed on the birds. Capturing flies off

exposed turkeys first involved immobilizing the host. Masking tape

was wrapped around the turkey's legs just above the feet which were

thus held together. The bird was placed on its side and a layer of

gauze followed by a strip of wide masking tape was placed across the

turkey's neck, chest, and legs and affixed to the platform which was

usually a large plastic tray. The upper wing was propped up to provide

the black flies with easy access to bare skin. Using this technique it

was possible to monitor the number and type of ornithophilic adults

present and capture them as well as observe their feeding locations,

duration and behavior. Their feeding could also be assisted and securing

of blood-fed flies from exposed infected turkeys could be more readily

ensured. To do this, flies that landed on the feathers of the wing or





-50-


elsewhere were quickly aspirated before they could scurry beneath the

feathers and were placed in the glass aspirator tube against the bare

skin of the underside of the wing elbow, the bare skin of the chest

over the rib cage or against the turkey's neck and allowed to feed to

repletion. The fly was observed, the feeding timed, and the turkey

soothed and kept as still as possible to assure a complete blood meal.

Once fed, the flies were transferred to paper pint ice cream cartons with

netting tops, provided a moistened paper towel to maintain high humidity

and transported to the lab. The exposed turkey technique was used to

collect black flies on numerous occasions from January through August

at 8 locations in 3 counties (Table 8).

To obtain black flies that had not been exposed to infections in

the field and to obtain specimens of known identity, especially when

working with species in the Simulium subgenus Phosterodoros which are

difficult to distinguish as adults and are much easier to separate as

pupae, adults were reared in the lab. Some adults were reared in pupal

vials from field collected pupae and other adults were reared form

field collected larvae and pupae in an aquarium with aerated water and

a fine mesh netting cover.

Black flies which fed on exposed infected hosts in the field were

held in the lab for at least two days after feeding and then allowed to

bite an uninfected bird. All adult flies maintained in the lab were

held in paper pint-sized ice cream cartons covered with netting and

provided with moist cotton for water and a raisin for nutrition. Up to

four cartons were kept at one time together in either a large plastic-

covered aquarium or a glass desiccator-type container (Fig. 6) where

high humidity was maintained by including a sponge half immersed in a



















































Figure 6. Glass container and paper cartons used for holding black
fly adults alive in the laboratory.









cup of water. Flies captured in an unfed condition in the field or

reared in the lab were allowed to feed on an infected turkey, held for

two to seven days and then allowed to feed on a clean bird. All

laboratory feedings involving uninfected hosts were conducted in a fly-

proofed former bull room at the Veterinary Science area at the University

of Plorida. This room was incandescent lighted,had a double door entrance

and had windows covered with very fine mesh organza material. Feedings

on infected hosts were conducted either in the Veterinary Science room

or at the fluorescent-lighted Veterinary Entomology Lab on campus.

Black flies were routinely taken off their sugar source at least 24

hours before planned feeding trials and maintained only on the water

provided by the moist cotton. Black flies were aspirated from the car-

tons or chilled in a refrigerator and removed with forceps and placed,

usually ane per vial, into seven-dram, clear plastic, cylindrical vials

covered with green, fine mesh (30/6.45 sq cm) netting at both ends and

held against the bare skin of the imrmobilized turkey. Room temperature,

relative humidity and feeding times were recorded.

Uninfected turkeys bitten by possible vectors were held in a fly-

proof room and the appearance of gametocytes in the peripheral blood

was monitored by blood smears obtained by wing vein punctures every 3

to 4 days. Blood smears were processed by air drying, fixing in

absolute methanol and staining in a 1:50 dilution of Giemsa solution

and distilled water. Flies were dissected in .9% physiological saline

and midguts and salivary glands were observed for the presence of

oocysts and sporozoites of L. smith. A few midguts were fixed in 10%

formalin, stained in Haematoxylin and mounted in balsam, with the

necessary intermediate steps for dehydration and clearing, in attempts





-53-



to more readily view the oocysts. Most midguts were air dried, fixed

in methanol and stained with Giemsa, as were the salivary glands. The

preparations were covered with permount and a cover slip for microscopic

examination. Other fly structures were placed in 10% Na0H (except the

wings) for 48 hours, rinsed in water and acid alcohol, placed in 100%

cellosolve for 15 to 30 minutes and mounted in balsam on a microscope

slide.














CHAPTER IV
RESULTS AND DISCUSSION



Simuliidae


General Comments


The primary efforts of this survey and ecological investigation

of the black flies of Florida were concentrated on the immature stages.

Over 700 adults were reared and pinned or mounted in balsam on slides

and a number of adults have been captured in traps or on host animals

(Tables 4, 6, 7 and 8) however the vast majority of specimens are

alcohol -preserved larvae and pupae. Over 1,100 four-dram vials con-

taining about 50,000 specimens from 1,080 individual, positive collections

which have been examined and identified are on hand. In addition,

records were obtained for Florida black flies from institution and

individual collections and are included under the appropriate species.

Table 1 is a list of the Simuliidae collected during this research

in Florida. Eighteen species representing two genera and six subgenera

are recorded. Figure 7 shows the 192 locations where I personally col-

lected specimens or obtained records from the collections of other indi-

viduals. One star normally designates a single collection site; in a

few cases a single star is used to indicate two collection sites adjacent

to each other such as a main stream and a side drainage flow to that

main stream. Each collection location is listed by county and site

number in the Appendix and the name and location of each stream is given.

The 50 counties from which black flies are recorded are listed in Table 2


-54-








Table 1. Florida black fly species.


Onephia (Chephia) ornithophilia Davies, Peterson, and Wood*

SimuZium (Byssodon) meridionate Riley

SimuZium (Byssodon) slossonae Dyar and Shannon

SimuZium (Eusimuiwnw congareenamrw(Dyar and Shannon)

SimuZium (Phosterodoros) diziense Stone and Snoddy*

SimuZium (Phosterodor~os) haysi Stone and Snoddy*

Sinulium (Ph~osterodoros) jenningsi Malloch

Simulium (Phosterodoros) jonesi Stone and Snoddy

Simutiurm (Phosterodoros) Zakei Snoddy*

Simulium (Phosterodoros) notiate Stone and Snoddy*

S~imultiwn (Phosterodoros) nyssa Stone and Snoddy

SiimuZium (Phosterodoros) tacodium Snoddy and Beshear*

SimuZiwn (Psitozia) vittatum Zetterstedt*

Simutium (Simuliwn) decorum Walker

SimuZiwon (Simutium) tuberosum (Lundstriim)

SimuZiwn (Simulium) verecundum Stone and Jamnback*

Onephia species undetermined No. l*

Simutium species undetermined No. 1*


New collection record for Florida.


































































Figure 7. Locations in Florida where black flies have been collected.










Table 2. Florida black fly distribution records by county.






Species



O O







County~~~~~~~ Sie

Alacua 3 x xx x x xx x x
Baker~Q 1 xx





Dade 1 x


Desoto
Dixie
Duval
Escambia
Flagler
Franklin
Gadsden
Glades
Gulf
Hamilton
Hardee
Hendry
Highlands
Hillsborough
Holmes
Jackson
Jefferson
Lafayette
Lake
Leon
Levy
Liberty
Madison
Manatee
Marion


x x
x x
x


x
x x
x
x
x x
x
x
X
x



x
x
x
x x
X
x
x
x
x
x x
x
x


X x
x xx
x x


x
x X


xX xx


x x x xx x
xX xx


x x x
x
x x


xX xx
xX xx


x
x xx


x xx
x x


x xx
x x x


x x
x
x


x x
X


x x xx
x x xX xx
x x X
x x xX x xx
xX x xx
x













Species


o 0








Osceola 1


Pinlla 1\ x



Sute 1 x x


Waul 1 xO~rO 3 ~


Wralng 4 x x xx


-58-


Table 2. (Continued)





























































By reading across or down from a selected species the number of times
the immatures of that species were collected together with the immatures
of another species is indicated by the figure at the intersection of the
two columns.


-59-


Table 3. Black fly associations based on collections of immature stages.*


26 1 4 4


84 3 1 38107 73
2 19 7

3 24
3 5
2 19 92
19 24 5 19 36


C. ornishophilia
S. meridionate
S. s~ossonae

S. congareenatum
S. diziense

S. haysi

S. jenningsi
S. jonesi
S. lakei
S. notiate

S. nyssa
S. taxodian
S. vittatum
S. decorum
S. tuberosum
S. verecundum

S. slpp. undetermined


2 2 32 10


52 15 186 55
6 2 49 23
21 4


73 4 21 8

2 1 40 11 3 253 44
150 15 54 19
23 1


7


38
1 107
73


92 36


8 44 10
8 9 50 33
9 67
4 50 6 78


2
2


32
10
No. 1


52 6 73 40 150
15 2 4 11 15


186 49 21 5 21 253 54
55 23 4 8 44 19


10 33


1 4


7 78


ar
s N


s s ~o
E~s
oEm


32





-60-


Table 4. Black flies captured in Manitoba traps.


Collection
times


1730-2030


1130-1930


1800-2000


1710-2020


1500-1745


0630-0930;
1730-2045

1630-1845


1600-1800


1600-1730


1500-1830


1530-1815


0700-1010;
1530-1800

1730-1845


0830-1430


1630-1805


1445-1710


Species
(all females)


vittartum


slossonae,
(Phosterodoros) spp.

s lossonae


slossonae


s lossonace


alossonae,
(Phosterodioros) spp.

slossnnae,
(Phosterodoros) spp.

s lossonae,
(Phosterodoros) spp.

s Zossonae


sZoss~nate


slossonae


stossonae,
(Phosterodoros) spp.

slossonae,
(Phosterodoros) spp.

slossonae


s lossonae,
(Phosterodoros)6 spp.

congrelcrenrum,
slossonae,
(Phosterodoros) CFpp


Date


1974: 3 May


30 May


28 June


28 June


6 July


11-12 July


16 July


16 July


19 July


19 July


24 July


26-27 July


15 Aug


17 Aug


15 Sep


1975: 24 Jan


Location


Univ. of Florida,
Site 34

Hatchet Creek
Preserve, Site 24

Lochloosa Creek,
Site 22

Hatchet Creek
Preserve, Site 24

Junction SR 225/
340, Site 8

Fisheating Creek,
Site 95

Lochloosa Creek,
Site 21

Hatchet Creek
Preserve, Site 24

Sandy Hatchet
Creek, Site 1

Univ. of Florida,
Site 34

Hatchet Creek
Preserve, Site 24

Fisheating Creek,
Site 95

Lochloosa Creek,
Site 22

Univ. of Florida,
Site 34

Lochloosa Creek,
Site ?22

Lochloosa Creek,
Site 22










Table 4. (Continued)


-61-


Collection
times


1000-1330


1100-1330


1000-1400



0930-1330


1030-1300


0930-1200


1400-1930


0730-1145;
1600-1910

1730-2000


0730-1200


0700-0830;
1500-1900

0830-1130;
1515-1715

0815-1315;
1620-2000

1625-1800


1400-1530


Species
(all females)


congareenarum,
s lossonae

congareenerumn,
s lossonae

s lossonae



slossonae


s lossonae,
(Phosterodoros) spp.

slossonae


s lossonae,
(Phosterodoros) spp.

sZossonae,
(Phosterodoros) spp.

stossonale


slossonae,
(Phosterodoros) spp.

slossonae,
(Phosterodoros) spp.

slossonae,
(Phosterodoros) spp.

slossonae


congareenarum,
slossonae

congareenrumro,
slossonae


Date


1975: 26 Jan


12 Mar


12 Mar


Location


Double Run Creek,
Site 43

Thomas Creek,
Site 159

8 km west of
Highway 115C on
Rt 90, Site 72

Double Run Creek,
Site 43

Turkey Creek,
Site 216

Junction SR 225/
340, Site 8

Hatchet Creek
Preserve, Site 24

Fisheating Creek,
Site 95

Lochloosa Creek,
Site 22

Fisheating Creek,
Site 95

Fisheating Creek,
Site 95

Fisheating Creek,
Site 95

Fisheating Creek,
Site 95

Turkey Creek,
Site 216

Double Run Creek,
Site 43


6


6


1


18


27-29


7


29-30


9-10


17-18


15-16


1976: 17


24













Collection Species
Date Location times (all females)


1976: 24 Mar Turkey Creek, 1640-1830 S. congareenzarum,
Site 216 S. slossonae

26 Mar Sante Fe College, 1030-1800 S. tuberosum,
Site 28 S. slossonae


-62-


Table 4. (Continued)












C. (C.)I ornithophilia

S. (B. ) meridionate

S. (B.)I slossonae

S. (E.)i congareenarum

S. (P.)I didiense

S. (P.)I haysi
------~I
S. (P.) jenningsi

S. (P.) jonesi
1 I
S. (P.) Lakei
I
S. (P.) notiate
I I

S. (P.) nssa

S. (P. ) taxodium

S. (P.) vittatum

S. (.S.)I decorum

S. (S.) tuberocum1

S. (S.)I verecundum
t
C. spp. No. 1

S. spp. No. 1


A S' O N D


J F M A M J J

Months


Figure 8. Seasonal occurrence of black flies in Florida.








with an indication of the species found in each county. Table 3 pre-

sents information on the frequency of association of black flies in

Florida's streams and rivers. Figure 8 shows the seasonal occurrence

patterns of the collected Simuliidae. Table 4 presents information on

species captured in Manitoba traps. Further reference to these figures

and tables will be made in discussing the individual species below.

Keys to the black flies of Florida, in all stages, are included.



Introduction to the Black Fly Keys


Characters which are important taxonomically on black fly larvae

and are noted on Figures 9, 10, and 11 include: the anterior, multi-

ple-rayed cephalic fans and their stalks (CF,CFS); the antennae (A); the

central dorsal section of the larval head capsule referred to as the

fronroclypeus or the cephalic apotome (CA) which bears usually dark

pigmented areas called head spots (HS); the ventral, setae bearing and

anteriorly toothed hypostomium or submentum (S); posterior to the sub-

mentum is the throat (or postgenal) cleft (TC) or gular notch which

differs in size and shape in the different species; laterally on the

thoracic region are a pair of organs called the respiratory histablasts (RH)

consisting of a number of coiled filaments whose number and branching

pattern are often diagnostic; the ventral proleg (VP); dorsally at the

posterior end the anal gills (AG), asmoregulatory organs, protrude and

may be simple and branlchless stalks or may have many branches and be

arborescent; behind the anal gills is located the anal sclerite (AS) or

X-piece which assists the larva in releasing its posterior from the silk

attachment patches; th7e cir-lfe or many rows of tiny anal hooks (AH) at




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CF r A
CFS CA















Figure 9. Dorsum of the head capsule of a black fly larva (S. slossonae).















TC


Figure 10. Venter of the head capsule of a black fly larva
(C. ornithophi. iia) .





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RO \


.*


Figure 12. Black fly pupa and cocoon (S. dixiense).


AS

Figure 11. Lateral view of two black fly larvae (S. dixiense).









the very end of the larva; and the ventral protrusions which occur on

some larvae and are called ventral or anal tubercles (VT).

Characters important on the pupa include: the filamented repira-

tory organs (RO); tiny setae-like structures on the thorax called

trichomes and posterior dorsal tail hooks (Fig. 12). On the

cocoon lateral apertures (LA) or windows occur anteriorly in the

Phosterodcros species and the texture and general regularity of the

cocoon is sometimes useful in separating species.

On the adult wing the presence or absence of hairs under the sub-

costal vein (SC); hairs on the dorsal, basal portion of the radius (R);

the presence or absence of a basal cell (BC); and color of hairs on the

stem vein (SV) are valuable characters (Fig. 13).

On the adult female head the shininess or pollinosity of the frons

(F) and its shape and that of the clypeus (C), the color of the antennal

segments (A), and the shape or size of the sensory vescicle of the

maxillary palps (SV) are useful taxonomically (Fig. 14).

Important characters of the male genitalia include: the appearance

of the ventral plate (VP) and the shape of its median portion; the shape

and relative lengths of the basimere (B) and distimere or clasper (D);

the presence or absence of a basal lobe on the distimere; and the number of

distal spines (DS) or teeth on the distimere (Fig. 15).

On the female hind leg the presence or absence of a dorsal groove

called the pedisulcus (P) on the second tarsal segment and the size of

the calcipala (CL), a flattened lobe on the inner side and at the apex

of the basitarsad segment, as well as presence or absence of a basal tooth

(BT) on the tarsal claws are important characters (Fig. 16j).

Structures useful for determinations in the region of the female





































Figure 13. A wing of the black fly Cnephia ornishophilia.


*


Figure 14. A frontal view of the head of a female black fly (S, notiaele.






























Figure 15. The male genitalia of a black fly, onephia omithophilia.







rC gBTI


Figure 16. The distal portion of the hind leg of a S. meridionatle
female.




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v g


Figure 17. The terminalia of a female S. meridionate.





-71-


genitalia (Fig. 17) are the size and shape of the genital. fork stem

(CF~S) an1d arms (CFA), the ovipositor lobes, each anal lobe (AZL) and

eachi cercus (CR).

A key to ther larvae of the blacki flies of: Florida.

(Photographiic illustrations of certain structures referred to in this

key and the keys which follow are included in the sections on the in-

dividual species. The larval key is primarily of value for later in-

stars. For further illustrations; refer to Stone and Jamnlhackk (1955),

Davies e~t al. (1962), Dood et al. (1963), StonE (1964), Stone and

Snoddy (1969~), Snoddy and Beshecar (1968) and Snotddy (11971, 1.976).

la. Hypostol:iuim convex, along anterior margin; head sp~ot pattern as in

Fig. 18. . .. ..... .. .. .. .. . C. ornrithop~hilia

16. Hypostomiumum cone~a\ or leve~l alo~ng tol anterio~r maergin~; the hcad

spot pattern not as in Fig. 18 ... .. .. ... . ... 2


2a. (Ib) Hiead spots light (white) .. .. .. .. . ... . 3

2b. Nlead spots dark or indistinct in a fulv~ous pattern . ... .. 5


3a. (2a) He~ad s5pot:, conpsristing of a cenitral.1 poste-io~r w~hte spot1 with

dark rays projecting and dive~rginge anterio~rly (Fig. 247, Stone and

Snoddy, 1.969; nol mature larva!e coll~ected~). . ... .S. mier~c~'idone

3b. Hea:d spot! s with! 3nterjior anld po~ster~ior- medial anld lalteral spots all

whiLe, spots vi-ib~le in thie usual pofitjions . .. .. .. .. 4


4a. (3b,) Meiccll an~terior and postecrior wh~ite hanld spots bor-dered by a

darki fulvous area on each sidle (F'ig. 99); Iespiratory his8tchlost

wilhl 8 fijlownts~n .. . ,. .. .. . . . ,'. decoruim

Ab. Nol dist inc. dairki fulvous border b~y rcontra? l he spots; histoblnst

wJith 6 filamints; .... . .. . . . . S. :'''^wrecundu









5a. (2b) Gular notch in the-form of a shallow inverted -v; headlspots

as in F~ig. 121 ... .. .. .. ... . .. C. species No. I

5b. Gular notch rectanngular, sub-rcctangular or sagiittiform. . .. 6


6a. (Sb) Cul~ar nlotch broadly sagittiform, ext~eundin at least across 8

of the venter of the head capsule, as widle as long .. .. .. 7

6b. Cular notch not Ragittiformi or else longer than wide .. .. .13


7a. (6a) R~espiracory histabla~sts consisting of 10 filomntins 9 of which

arise fr~om a thick basal 10th (Fig. 60); aaterior medial head spots

often weak . .. ... .. .. ... .. .. S. Jonesi

7b. Respiratory histablast not as above; a.ll head spots usually dark,

distinct .. .. .. ... .. .. .. .. .. .. ... 8



8a. (7b) Respiratory histroblast with 6 filaments; anal tubercles not

prominent. .. .. .. .. .. .. ... .. . .. S. notiate

8b. Respiratory histoblast within more than 6 filarmen~ts; anal tubercles

promninenlt........ ... ... .... ........9


9a. (8b) Respiratory histoblast wait-h 7 filanones. . . .. S. haysi

9b. Respiratory histablast with more than 7 filame-ncs. .. .. .10


10a. (9b) Respiratory hristoblast with, 8 filaments. . ... S. tiaOicodiim

10b. Respiratory histohlast with more than 8 filamnents. . ... .11


11a. (10b) Rlespiratory histablast ulith 9 ;ilanments .. .. .. ,S. Zakei

11~b. Respiratorq histohlzst wJith 10 finments . .. .. .. .12


1.2a. (1lb) 10 recspira::ory h-iotablast filamennts w~ith a pattern as in

Fig. 81.; a1nal tuberele~s piromuinenl 91nd CocaiI. .. . S. niYouC

1.2b. 10 respiratory hiistchlast filametnts w~ith a palttorn as in Fic. 53;

anal tuibercler sma1 :l and ronded lc.. . .. . .. jennringsjri










13a. (6b) Gular notch long extending b the distance or more to thle teeth

of the submentum, sagittiform or subrectangular, pointed or rounded

anteriorly. .. ... .. ... .. ... . . . 14

13b. Gular notch extending less than k the distance to the teeth of sub-

mentum, subrectangular. ....... ...... .. 16


14a. (13a) H-ead spots indistinct in fulvous area; gular notch usually

with parallel sides; anal tubercles inconspicuous . S. tuberosure

14b. Head spots distinct; gular notch with curved margins, elongate

sagittiform; anal tubercles prominent . . . . . . 15


15a. (14b) Re~spiratory histablast with 6 filaments; larvae reddish.

. . . . . * S. slossonae

15b. Respiratory histoblast with 10 filamren~ts; larvae ye~llow.

. . . . . . . . . . - S, clixience



16a. (13b) Respiratory histoblast filaments 4 in number; gular notch

longer than wide. . .. ... . .. . . S. species No. 1

16b. Respiratory histoblast. filaments numbeLr more than 4; the gular

notch not longer than wide. .. .. .. .. ... . .. .. 17


17a. (16b) Respiratory histablast with 12 filaments; the anal tubercles

are large, prominent; a medium~-sized larva, 6 mm~ long.

. . . S. conga~ailreeam

17b. Respiratory histoblast withi 16 filaments; the anal tubercles are

absent or inconspicuous; a large larva, 8-9 mml long . S. viCttatumi






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A key to the pulPae of the black flies of Florida.

(No pupae of Cnephiia species No. 1 have been observed.)

la. Cocoon a loose mass of silk, indistinct shape; strong dorsal books

at the posterior end of the pupa. . . .. .. C. ornithophilia

lb. Cocoon a distinct slipper or pocket shape, strong dorsal hooks

absent. .. .. .. .. .. .. .. .. .; . . .. . 2


2a. (lb) Cocoon with large anterior, lateral apertures .

2b. Cocoon uniform without large lateral apertures. ...



3a. (2a) Pup~a wath 6 respiratory filaments on each side;

antero-ventrally comlpletely joined (FiG. 75). ....

3b. Pupa with miore than 6 filaments ...........



4a. (3b) Pupa with 6 filaments rising off a strong basal

(Fig. 48) . . . . . . . . . . .

Ab. Pupa with more tha7n 7 fil.aments ...........



Sa. (4b) Pupa with 8 filamecnts with four pairs of 2. ..

5b. Pupa wit-h more than 8 fi'l~anclts ..........


. . . 3

. . .10


cocoonI

..S. notiate





7h one

...S. haysi





.S. Zaxodium

. . .6


6a. (5b) Pupa with 9 filaments in the pattern 2, 2, 2 and 3 from the

dorsal. .. .. ... .. .. .. .. .. .. .. .. S. lakei

6b. Pup3 with 10 filaments. ... .. .. ... .. .. .. 7



7;1. (6b) Pupa wJithl 9 filaments rising from a thick basal 10h one

(Fig. 60) ... ... . .. ... . ... .. ionesi

7b. Pupal respiratory organ lacking a str-ong basal filnment . .. 8



88. (7b) Pupa w~itn 10 respiraltoryy filamecnts whichh rise as 5 pairs, the

petiole for filaments 7 nnd 8 (from; thIe dorcal) noticiably longer










and stouter than the other petioles (Fig. 12) .. .. S. diziense

8b. Pupa with 10 filaments, some of which~ rise in pairs others in

triplets. .. .. .. .. . .. . .. . .. 9



9a. (8b) The lower more ventral respiratory filaments on long petioles,

filaments long. .. ... ... .. .. .. .. .. S. nyssa

9b. Lower filaments rise close to the base, petioles short, filaments

short .. .. .. ... ... .. .. . S. jcinnngsi



10a. (2b) Pupa with 4 respiratory filaments. .. .. S. species No. 1

10b. Pupa with more than 4 filaments .. .. ... .. .. 11



11a. (10b) Pupae with 6 filaments in the respiratory organ1. .. 12

11b. Pupae with more than 6 filaments. . . ... . 14



12a. (11a) Pupal cocoon with concave anterior edge (in lateral view);

respiratory filaments rising from long petioles (Fig. 28)S. slossonae

12b. Pupal cocoon with convex or straight anterior edges; petioles

short ... .. .. .. ... . .. .. . ... .. .3



13a. (12b) Respicatory filaments widespread, base of filamlents 5 and

6 sharply separated from banse of 3 and 4,filaments long (Fig.115).

. . . . . . * * S. vercun dw~n

13b. Respiratory filaments not: widespread, bases about equidistant,

filaments short (Fig. 108). ... .. .. ... .. S. :ubcr'oswni



14a. (11h) Pupa with 8 respiratory f~laments; cocoon with rouGh text~ure.

. . . . . . . . . . S. accr:C r nl

14b. More than 8 filaments . .. .. . .. .. .. . 15



.15a. (141,) Pupa with 12 respiratory filaments; cocoon with long anterior-





-76-


dorsal projection (Fig. 35).. . .. ... .. .. cong'areenarurr

15b. Pupa with more than 12 filaments; cocoon lacking dorsal projec-

tion .. .. .. .. .. .. .. .. . .. . .. .. 6



16a. (15b) Pupa with 16 filaments in 8 pairs . .. .. .. S. vittatus

16b. Pup3 withr more than 20 filaments in pairs and in 3's .S. meridionale




A key to the adult male black flies of Florida.

(Adapted and modified from Stone and Enoddy, 1969)

la. Second tarsal. segment of the hind leg without a pedi~sllcus but a

shallow~ depression may be present; radius vein with hair~ dorsally

on the basal segment; large basal cell present . .C. ornithophilia

lb. Second tarsal sfement of the~ hind leg with a deep pedisulous; base

of radius withi or without hair dorsally; large basal cell absent.





2a, (Ib) Basal portion of radius with hair dorsally S. congareeniar~um

2b. Basal. portion of radius without hair dorsnlly. .. .. .. .. 3


3a. (2b) D~istimere~ short, stout, with; 3 or mo~re tceeth .. S. visttumw

3b. Distimore long, with 1 tooth or none .. .. ... .. .. 4


4a. (3b) Distimere with a rounded 10be on the inner margin near the

base .. . . .. .... .. ... .. ... .. 5

4h, Distimefre without a lobe or basal projection .. .. .. .. 6


Sa. (42) Distimuere with ba~sal lobe bearing a number of stout, spine-

Itke scenec . .. .... .. .. .. ... .. S. tuber'osur

5b. Distime~re with basal lobe bearing finec hairs only. . S. stossonae










6a. (4b) Ventral plate In ventral view broadly rounded. S. meridionale

6b. Ventral plate in ventral view more narrow, compressed from th~e

sides. . .. . ... . .. .. .. . .. .. 7



7a. (6b) Ventral plate with basal arms hearing distinct lateral pro-

jactions; posterior third of sc~utum shiny with indistinct hairs. 8

7b. Ventral plate with basal arms without distinct lateral projections;

posterior quarter or less of scutum shiny with some strong erect

hairs. ... .. .. ... . .. ... .. .. . .. 11



8a. (7n) Ventral plate in ventral view with median portion longer than

wide and parallel-sided or nearly so; scutumn with silver spots

narrow, oblique; thle dark area be~tween the spots broadens anteriorly.

. . . . . . . . . . . . . . S. dizience

S. jenningjsi

S. notiate


8b. Ventral plate in ventral view with median portion not longer than

wide and' widened toward the end; anterior silver spots ;nd inter-

vening dark area variabLe. .. .. ... .. .. ... .. 9



9a. (8b) Scutuml with large anterior silvery spotsa each extendingi about

one-third the distance across the front of thle scutum; thle dark area

between the spots na1rrows strongly to thle anterior margin. .S. Jone~d

S. txodiuim


9b. Scutumi with smanller silvery spots; ilhe dark area betweenr th:e spots

is broad convergies little, and is narrow~est a-bout mindway along

the l~ength of the spot; .. .. .. .. ... . S. hiaUGL

S. lakei.

S. nuiscal









10a. (7b) Ventral plate in ventral view narrow, V-shaped, middle

region tapers almost to a1 point; with a1 ventral keel. S. decorumr

10b. Ventral plate broader, not as strongly compressed in the middle

section; ventral keel absent. .. .. .. .. .. S. vereoundlun




A key to the adult female black flies of Florida.

(Adapted and modified from Stone and Snoddy, 1969)

la. The second hind tarsal segment lacks a pedisulcus, although there

may be a slight depression; the basal cell is present and the basal

portion of the radius bears setae dorsally . .. C. ornlithophiliai

lb. The second hinid tarsal segment with a deep pedisuicuis; large basal

cell absent and the basal portion of the radius bears or lacks setae

dorsally (Simu~7~ium.. ... ... .... . . . . ..


2a. (lb) Radius vein with hair dorsal~ly on the basal section.

. . . .. . . . .. S congareelnarumn

2b. Radius without hair dorsally on the babal section. .. .. .. 3


3a. (2b) Tcrsal claw with a prominent basal. tGOoc .. .. .. .. 4

3b. Tarsal claw simple, without a prominent basal tooth. .. .. 5


4a. (3a) Fr-ons gray pollinose .. ... .. .. .. S. mieridionale

Ab. Frons shiny black. .. .. ... .. .. ... .S. slossonna


Sa. (3b) Frons and terminal abdominal targites shiiny black or dark

brown. ... . .. .. . . .. ... . .. . .. 6

5b. Fron~s and tecrminal abdominal tergites at least lightly pollinose.10


6a. (Sa) Sub~cosr-al vein with a row of hai~rs ventral3y; scutumn

subshiny .. .. .. .. ... . . .. . .. .7










6b. Subcostal vein without hairs ventrally; scutumi shiny .. .. 8


7a. (6a) Fore tibia with a narrow gray-white patch not more than one

third the width of the tibia; inner margins of ovipositor lobes

fairly st-raigiht. .. .. .. .. .. .. .. ... S. tubhero~swn

7b. Fore tibia with a brilliant white patch which covers one half or

more of the tibia; inner margins of the ovipositor lobes concave

enclosing anr oval area .. .. ... ... .. S. ver~ecundum


8a. (6b) In anterior view the scutum displays a pair of fairly distinct

rounded pollinose spots with a darker area of the scutum in between;

hairs on the stem vein are dark brown to black . .. S. jenningsi

S. notiate

S. nyssa

S. taxodium

8b. In anterior view the scutum is not pollinose or the pollinosity

occurs as a diffuse area along the front and sides .. .. .. 9


9a. (8b) Clypeus about as wide as long. . .. .. S. jonesi

S. Zakei

9b. Clypeus longer than wide . .. . .. .. .. S. dixiense

S. haysi


10a. (5b) Scutum silvery gray with dark brown markings; abdomen with a

black and light gray pattern .. .. .. .. .. S. v~ittatusm

10b. Scutum uniform brownish gray without contrasting dark brown manrkings;

abdomen black with thin g~ray pollinosity without a pattern.

. . . . . . . . . . S. decolmm





-80-


Introductionto th Individual Species Sections


The followingf sections deal specifically with the black fly species

found in Florida. References are listed below each species name which

provide synonyms for the currently accepted name and additional sources

of descriptions andi figures. The descriptions which follow the brief

taxonomiy portions are intended to point out outstanding or diagnostic

characters and are accomlpanied with photographic illustrations. Under

"Florida Observations" below the heading "Stream Parameters" A summary

is given of the dimension, pH, temperature, and velr.city figures for

streams which contained each par-ticular species. i~n the sectionn on

Florida collection records the numbers i:amediatel~y inl!.owing the county

names refer to individual collection siteS identified inl the A~ppendix.

The code for thle collection records is as Follow~s: S refers to small,

very young larvae with no or weakly developed respiratory histoblasts;

M refers to medium-aged larvae with distjinct but white histablasts;

Refers to large, mature larvae withi dar:k histobl~asts; P refers to

full pupae or pupal exuviae; C refers to ucocons; anld A1 refer~s to male

or female adults. Tlh- majority of records refer to specimens the

authonr personally collected; in those instances where specimens were

gathecred by other individuals, the collector is noted. On the indi-

vidual species ma~ps wJhen a record is included with the only locality

information beingi the county the mark for that record on the ma~p is

placed in thle center of the :ounty. Immat~nure, very young larvae

lacking well-develouped respirnatry h~istobtasts in the subgenus

Phol~sterodoros~? are difficult if not impossible to differentiate as: are

the adlul~t fema;les. Records for t~he P~hostr:: odo:. l~.i ronpeces are based~

prima~rly oin mature larvae and p~uPre. Only at locntionss whero









essentially a single Phosterodoros species was found to exist are

records for earlier stages included. Records for undetermined species

are listed as S. (Phosterodoros) spp. in the Manitoba trap results

(Table 4). Dr. E.L. Snoddy kindly examined immature and adult specimens

of S. jenningsi, S. lakei and S. tarodium~ from Florida and confirmed my

identifications.



Onephia (Cnephia) or-nithophilia Davies, Peterson, and Wood


onephia ornitshophiziai Davies, Peterson and Wood, 1962, Proc. Entomol.

Soc. Ontario 92: 102 (female).

Onephzia ornithoph2iia Stone and Snoddy, 1969, Auburn Univ. Agr. Exp.

Sta. Bull. 390: 25 (female, larva).


Taxonomy. Davies et al. (1962) first described a female of this

species. The holotype was collected from a blue jay at Algonquin Park,

Ontario and was deposited in the Canadian National Museum. Paratypes

are located in the U.S. National Museum. Stone and Snoddy (1969) in-

dicated the male of C. orn)ithophlZ~ia was not known and that the larva

and pupa were apparently indistinguishable from those of C. pecuaruo.

Description. The larva is large, about 8 mm long with a grayish

brown abdomen and a light yellow brown head capsule. The cephalic

spots are dark with the median groups of about 20 forming one long

continuous row (Fig. 18). The submentum is convex apically and bears

small teeth. The gular notch has fairly parallel sides, is broadly

rounded anteriorly and extends less than 1/3 the distance to the sub-

mental teeth (Fig. 10). The cephalic fans each contain about 60 rays

with long hair-like spines. The anal tubercles are absent.





-82-


Figure 18. The head spots of a larva of C. ornithophilia.


Figure 19. The pupal exuvium and cocoon of C. ornithop~hilia.





-83-


The pupa bears strong dorsal posterior hooks and is located in an

irregular, loosely woven cocoon. The respiratory organs each contain

about 30 filaments (Fig. 19).

This is believed to be the first description of the male. The

wings are 3.75-4 mm long. There are hairs dorsally on the base of the

radius. The first segment of the flagellum is the longest, about twice

the length of the other segments. The scutum is dull brownish black with

numerous short golden hairs. The scutellum is reddish brown and bears

long dark hairs. The legs are almost uniform yellow brown. The second

hind tarsal segment lacks a pedisulcus. The abdomen is dull brownish

black dorsally and lighter gray or light brown ventrally. The ventral

plate in ventral view is broad and the tapering distimere which is

almost equal in length to the basimere has a single tooth at its apex

as in Fig. 15.

Plesiotype: Male, with associated pupal exuvium and cocoon,

NW 23rd Ave. and NW 83rd St., Gainesville, Alachua Co., Florida, 2

February 1975 (Pinkcovsky), to be deposited in the U.S. National Museum.

The wings of the female are about 4.5 mm long. The base of the

radius is covered with setae and a basal cell is present on the wing.

The frons bears short yellow hair. The pedisulcus is essentially absent

and the prominent tooth on the basal claw has convex margins (Fig. 20).

The apex of the abdomen is shiny. The genital fork has widespread

narrow arms as in Fig. 21.

Distribution. Stone and Snoddy (1969) indicate C. orni-thophiia

occurs in Louisiana, Mississippi, South Carolina, Virginia, and Ontario.

Tarshis and Stuht (1970) report this species from Mlaryland.

Life History. Stone and Snoddy (1969) mention that C'. ornith~ophilla







































Figure 20. Tarsal claw of a female of C. ornithophilia.


Figure 21. Genital fork and terminalia of a
female.


C. ornithophilia









is probably univoltine. Eggs collected in stream bottom samples and

placed in laboratory rearing tanks yielded first instar larvae three

days later (Tarshis, 1973). Larvae have been collected between 11

January and 18 March in South Carolina and adult females were reared on

6 February from pupae; females have been collected from 6 February to

1 May in Louisiana and Mississippi (Stone and Snoddy, 1969). Tarshis

(1973) working in Maryland found larvae from 14 November to 29 April,

pupae from 11 January to 22 April and adults from 2 to 17 March. Tarshis

and Stuht (1970) captured adult C. ornithophitia between 28 February

and 1 April in emergence cages over a stream in Maryland. When first

to third instar larvae were collected and reared in the laboratory in

aerated water at 15-220C pupae appeared after 2 to 21 days and adults

emerged 4 to 23 days after introduction of the larvae (Tarshis, 1973).

Tarshis (1973) found adults emerged as soon as 2.5 hours after pupation

[very difficult to believe] and that the number of males emerging ex-

ceeded the number of females emerging only on the first day.

Ecology. Tarshis and Stuht (1970) found massive numbers of larvae

on leaves, rocks, and twigs in a stream that flowed .153 to 1.0 m/sec

(.49-3.3 ft/sec) from a pond through an oak and willow woods. TIhe

stream was .92-1.53 m (3-5 ft) wide, 2.5-20 cm (1-8 in) deep and had a

pH reaction between 6.9 and 7.4. In a similar stream that was slightly

deeper and 1.22-3.66 m (4-12 ft) wide, larvae were collected at water

temperatures of .5-15oC (Tarshis, 1973). Tarshis (1973) mentioned that

pupae easily suffered injury and that development to adults did not

occur when pupae were removed from their substrate. Larvae of C. muitaba,

P. gibsoni, S. decorum, S. tuberosumn, S. venustum, and S. vitt~atum were

collected with C. ornithophilia in a pond runoff stream where









C. ornithophilia predominated (Talrshis antd Stuh~t, 1970).

Habits. Tarshis and Stubt (1970) suggest that C. ornithophilia

ov-iposits on the stream surface and the eggs sink to the stream bed.

Bennett (1960) in Ontario recovered C. ornithophilia (as CnephiJa "U")

fromt jays, hawks, robins, sparrows and other woodland birds. Tarshis

(1972) fed C. onnithophiia on geese and ducklings in the laboratory

and transmitted L;. simondi to ducklings by the bite of the fly. Tarshis

(1973) did not collect any C. ornithophi~lia from ducklings exposed on

the banks of a stream containing larvae and pupae of the black fly and

suggests that thle target hosts were inappropriate or perhaps that the

flies were absent at ground level, and concentrated in the canopy.

Florida observations.

Stream Parametrers

Width Depth pH Temperature Velocity

Mleanl: 2.57m 23.64 cm 4.7 14.70C (58.60F) .45 m/sec (1.47 ft/sec)

Mlin: .3 1.3 3.6 8.3 (47) .23 (.75)

Max: 11 90 6.6 24.4 (76) 1 (3)


C'nephia ornithophilia-lc is reported here for the first timne from

Florida. Both Stone (19)65) and Stone end Snoddy (1969) mention that

C. pecuarum occurs in Florida. Key chiaracters, such ats the shape of

the tooth on the tarsal claw of the female, on specinens I obtained in

Florida differed from those of C. pccuarwn.m In the U.S. National

Museum I located one adult On~ephia specimen from Florida, listed for

Site 204 below,, which was a male that had been labeled as C. pecuarum

but C. ornithophiliadn was pe~nciled inl underneath.. Adult and immature

specimens whlichl I submitted fro~m five collections representingli two welst

Florida (Hol~mes and Liberty) and one east F~lorida (Alachua) counties werre








identified by G.E. Shewell of the Canadian Biosystematics Research

Institute as C. omnithophilia.

Onesphia ornithophilia was collected from 31 sites in 14 counties

in Florida (Fig. 22). Most of the records represent collections of only

a few, usually young larvae with small, pale histoblasts, but the larger

overall body size compared to other species and distinctive head spots

and gular notch are discernible even in young larvae. Larvae, pupae

and associated adults were obtained at three sites 28, 119 (Fig. 23),

and 147). This species was first collected in the streams on 28 October

and last collected on 18 April. Most collections occurred and the

largest populations were encountered during December through March.

Collections of C". ornithophilia at a number of sites were made during

one winter month and the species was not found again until about the

same time the following year. It appears that C. ornithophilia is

univoltine in Florida. This species occurs in cool, fairly shallow

flows, usually at most a few meters wide, most frequently with a velocity

of .305-.61 m/sec (1-2 ft/sec) and a pH which normally is below 5. Im-

matures were also collected at Lebanon Station (Site 139) and Yellow

Water Creek (Site 70) where the pH approaches neutrality. At most sites

there was considerable vegetation especially trailing grass in the stream

to which the immatures were attached. At Blues Creek (Site 2) and Site

28 sparce vegetation was present. Site 28 originated in a small swamp

just upstream, was about .5 m wide, 10 cm or less deep and tumbled down

a root-filled and rocky bed through woods. In January a heavy popula-

tion of C. ornithophilia covered rocks, dead tree leaves, pine needles,

and a few blades of trailing grass. Masses of 40 or more larvae en-

circled twigs about .6 cm in diameter in the current. Most of the




Full Text



PAGE 1

THE BLACK FLIES (D1PTERA: SIMDLIIDAE) OF FLORIDA AND THEIR INVOLVEMENT IN THE TRANSMISSION OF Leuoooytozoon smithi TO TURKEYS By DENNIS DREW PINKOVSKY A DISSERTATION PRESENTED TO THE GRADUATE COUNCIL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPH"* UNIVERSITY OF FLORIDA 1976

PAGE 2

ACKNOWLEDGEMENTS I sincerely thank Dr. J.F. Butler, my Committee Chairman, for his constructive suggestions and support during my research endeavors and for his valuable, critical review of my dissertation. To Dr. D.F. Forrester I extend my deep appreciation for the advice and guidance ha enthusiastically offered during my transmission studies and for the o.quipment and facilities he generously allowed me to use. To all the members of my Ph.D. Committee I express appreciation for their critical appraisal of my dissertation and their helpful comments . I wish to thank Dr. E.L. Snoddy and Dr. G.E. Shewell for examining black fly specimens which I sent from Florida and for the determinations rendered. I express my gratitude for the opportunity to examine black fly specimens collected in Florida which were made available to me by numerous individuals and institutions. To the staff at the U.S. Department of Agriculture Insects Affecting Man Laboratory, Gainesville, I extend my appreciation for the use of equipment and for advice during my research and academic studies. I am grateful to Dr. E.V. Komarek and the staff at the Tall Timbers Research Station and L.E. Williams, D.H. Austin, and T. Peoples of the Florida Game and Fresh Water Fish Commission for the use of facilities and other assistance during my collecting trips.

PAGE 3

I wish to also thank the State of Florida Division of Recreation and Parks for collecting permits and the opportunity to gather specimens at the beautiful State Parks around Florida. I extend my deep gratitude to the U.S. Air Force Institute of Technology Civilian Institutions Program for financial support and for the opportunity to attend the University of Florida and pursue my Ph.D. degree. To P. Humphrey, D. Young, L. DuBose and especially T. DiNuzzo I express my sincere thanks for good-spirited support during my research. iii

PAGE 4

TABLE OF CONTENTS Page ACKNOWLEDGEMENTS ii LIST OF TABLES • vi LIST OF FIGURES vii ABSTRACT xiii CHAPTER I. INTRODUCTION 1 II. LITERATURE REVIEW 3 Simuliidae 3 Taxonomy and Distribution 3 Bionomics 7 Damage 22 Control 23 Florida Ecological Habitats 27 Leuooaytozoon smithi 33 III. MATERIALS AND METHODS 40 Black Fly Survey 40 Leuaooytozoon smithi Transmission 45 IV. RESULTS AND DISCUSSION 54 Simuliidae 54 General Comments 54 Introduction to the Black Fly Keys 64 A key to the larvae of the black flies of Florida 71 A key to the pupae of the black flies of Florida 74 iv

PAGE 5

TABLE OF CONTENTS (Continued) Page A key to the adult male black files of Florida 76 A key to the adult female black flies of Florida 78 Introduction to the Individual Species Sections. . . 80 Cnephia (Cnephia) ornithophilia Davies, Peterson, and Wood 81 Simulium ( Byssodon) mevidiondle Riley 92 Simulium (Byssodon) slossonae Dyar and Shannon ... 99 Simulium (Eusimulium) congareenarum (Dyar and Shannon) 117 Simulium (Phosterodoros) dixiense Stone and Snoddy . 128 Simulium (Phosterodoros) haysi Stone and Snoddy. . . 137 Simulium (Phosterodoros) jenningsi Malloch 143 Simulium (Phosterodoros) jonesi Stone and Snoddy . . 155 Simulium (Phosterodoros) lakei Snoddy 169 Simulium (Phosterodoros) notiale Stone and Snoddy. . 181 Simulium (Phosterodoros) nyssa Stone and Snoddy. . . 189 Simulium (Phosterodoros ) taxodium Snoddy and Beshear 195 Simulium (Psilozia) vittatum Zetterstedt 206 Simulium (Simulium) decorum Walker 224 Simulium (Simulium) tuberosum (Lundstrcm) 235 Simulium (Simulium) verecundum Stone and Jamnback. . 255 Cnephia species Undetermined No. 1 268 Simulium species Undetermined No. 1 271 Leucocytozoon smithi Transmission 277 V. CONCLUSIONS 294 LITERATURE CITED 296 APPENDIX. COLLECTION SITE NAMES AND LOCATIONS 319 BIOCRAPHICAL SKETCH 331

PAGE 6

LIST OF TABLES Table Page 1. Florida black fly species 55 2. Florida black fly distribution records by county 57 3. Black fly associations based on collections of immature stages 59 4. Black flies captured in Manitoba traps 60 5. Sentinel turkey locations and results 278 6. Black flies captured in ramp traps 281 7. Blackout box trapping results 282 8. Black fly captures from exposed turkeys 283 9. Leucooytozoon smithi transmissions 289

PAGE 7

LIST OF FIGURES F1 S ure Page 1. A modified Manitoba trap with a black plastic skirt .... 44 2. Sentinel turkeys in an exposure cage 47 3. A blackout box trap in the field 47 4. One view of a ramp trap 48 5. An exposed turkey in the field 48 6. Glass container and paper cartons used for holding black fly adults alive in the laboratory 51 7. Locations in Florida where black flies have been collected 55 8. Seasonal occurrence of black flies in Florida 63 9. Dorsum of the head capsule of a black fly larva (S. slossonae) 65 10. Venter of the head capsule of a black fly larva (C. ornithophilia) 65 11. Lateral view of two black fly larvae (S. dixiense) 66 12. Black fly pupa and cocoon (S. dixiense) 66 13. A wing of the black fly Cnephia ornithophilia 68 14. A frontal view of the head of a female black fly (S. notiale) 68 15. The male genitalia of a black fly, Cnephia ornithophilia 69 16. The distal portion of the hind leg of a S. meridionale female t 69 17. The terminal i a of a female S. meridionale 70 18. The head spots of a larva of C. ornithophilia 82 vii

PAGE 8

LIST OF FIGURES (Continued) Fig^e Page 19. The pupal exuvium and cocoon of C. ornithophilia 82 20. Tarsal claw of a female of C. ornithophilia 84 21. Genital fork and terminalia of a C. ornithophilia female. . 84 22. Collection locations for C. ornithophilia in Florida. ... 88 23. Site 119, Gum Creek, a stream inhabited by C. ornithophilia 89 24. The pupa and cocoon of S. meridionale 94 25. The scutum of a S. meridionale female 94 26. Collection locations for S. meridionale in Florida 97 27. Gular notch of a S. slossonae larva 101 28. The pupa and cocoon of S. slossonae 101 29. Terminalia of a male of S. slossonae 102 30. Terminalia of a female of S. slossonae 102 31. Collection locations for S. slossonae in Florida. ..... 105 32. Site 43, Double Run Creek, where S. slossonae immatures were collected _ 107 33. Cephalic apotome of a S. congareenarwn larva 118 34. Venter of the larval head capsule of 5. congareenarwn . . .118 35. Pupa and cocoon of S. congareenarum 120 36. Terminalia of a male of S. congareenarum 120 37. Terminalia of a female of S. congareenarum 121 38. Collection locations for S. congareenarum in Florida. . . . 124 39. Site 216, Turkey Creek, a typical S. congareenarum stream 125 40. Dorsal view of the head capsule of a S. dixiense larva. . . 130 41. Gular notch and hypostomium of a S. dixiense larva 130

PAGE 9

LIST OF FIGURES (Continued) Figure Page 42. Male terminalia of S. dixiense 132 43. Terminalia of a female of S. dixiense 132 44. Collection locations for S. dixiense in Florida 134 45. Site 74, Pine Barrens Creek, a stream inhabited by S. dixiense 135 46. Cephalic apotome of a S. hay si larva 138 47. Gular notch of a S. hay si larva 138 48. Pupal exuvium and cocoon of S. hay si 139 49. Location of the collection site for S. haysi in Florida . . 141 50. Site 195, Juniper Creek, where S. haysi was collected . . . 142 51. Cephalic apotome of a S. jenningsi larva 145 52. Gular notch of a S. jenningsi larva 145 53. A pupa and cocoon of S. jenningsi 146 54. S. jenningsi male terminalia 146 55. Genitalia of a female of S. jenningsi 147 56. Collection locations for S. jenningsi in Florida 151 57. Site 141 at Gulf Hammock where S. jenningsi was collected . 152 58. Cephalic apotome of a S. jonesi larva 156 59. Gular notch of a S. jonesi larva 156 60. Respiratory organ of a S. jonesi pupa 157 61. Terminalia of a male of S. jonesi 159 62. Genital fork and terminalia of a female of S. jonesi. . . . 159 63. Florida collection locations for S. jonesi 161 64. Site 210, the Fenholloway River, a S. jonesi collection location 162

PAGE 10

LIST OF FIGURES (Continued) Figure Page 65. Cephalic apotome of a 5. lakei larva 171 66. Gular notch and hypostomiura of a S. lakei larva 171 67. Pupa and cocoon of 5. lakei . 172 68. Dorsal view of a male of 5. lakei 172 69. Terminalia of a male of S. lakei 173 70. Terminalia of a female of S. lakei 173 71. Collection locations for S. lakei in Florida 175 72. Site 135, Otter Creek, a collection site for S. lakei . . . 176 73. Head spots of a S. notiale larva 183 74. Gular notch of a S. notiale larva 183 75. Pupal exuvium and cocoon of S. notiale 184 76. Scutum of a male of S. notiale 184 77. Terminalia of a male of S. notiale 185 78. Terminalia of a female of S. notiale 185 79. Collection locations for 5. notiale in Florida 187 80. Site 88 at Chattahoochee where S. notiale immatures were found 188 81. S. nyssa pupa and cocoon 191 82. Collection locations for S. nyssa in Florida 193 83. Site 116, Blue Creek, where S. nyssa was collected 194 84. Cephalic apotome and head spots of a S. taxodium larva. . . 197 85. Gular notch of a S. taxodium larva. 197 86. Pupal exuvium and cocoon of S. taxodium 198 87. Male terminalia of S. taxodium 198 88. Female terminalia of S. taxodium 199

PAGE 11

LIST OF FIGURES (Continued) Figure Page 89. Collection locations for S. taxodium in Florida 201 90. The Ichetucknee River, a S. taxodium collection site. . . . 203 91. Head spots of a S. vittatwn larva 209 92. Gular notch of a S. vittatwn larva 209 93. 5. vittatwn pupa and cocoon 210 94. Terminalia of a S. vitiation male 210 95. Scutum of a female of S. vittatwn 211 96. Terminalia of a female of S. vittatwn 211 97. Collection locations for S. vittatwn in Florida 217 98. Site 167 at Crestview where S. vittatwn was collected . . . 219 99. Head spots of a S. decorum larva 226 100. Gular notch of a S. decorum larva . 226 101. 5. decorum pupa and cocoon 227 102. Male terminalia of S. decorum 227 103. Female terminalia of S. decorum 228 104. Collection locations for S. decorum in Florida 232 105. Shepard's Mill, Site 86, where S. decorum was collected . . 234 106. The cephalic apotome of a S. tuberosum larva 237 107. The gular notch of a S. tuberosum larva 237 108. Pupa and cocoon of S. tuberosum 238 109. Male terminalia of S. tuberosum 238 110. Terminalia of a female of S. tuberosum 239 111. Collection locations for S. tuberosum in Florida 243 112. Site 56 in Clay County, a collection spot for S. tuberosum. 2^6

PAGE 12

LIST OF FIGURES (Continued) Figure Page 113. The head spots of a S. vereaundum larva 257 114. The gular notch of a S. vereaundum larva 257 115. The pupa and cocoon of S. vereaundum 258 116. Male terminalia of S. vereaundum 258 117. Terminalia of a female of S. vereaundum 259 118. Collection locations for S. vereaundum in Florida 262 119. Panther Creek, Site 223, where S. vereaundum was collected .264 120. Cephalic apotome of a larva of Cnephia species No. 1 . . . .269 121. Venter of the head capsule of a larva of Cnephia speaies No. 1 269 122. Collection location for Cnephia species No. 1 in Florida . .270 123. Pupal exuvium and cocoon of Simulium speaies No. 1 273 124. Larval head spots of Simulium speaies No. 1 274 125. Venter of the head capsule of the larva of Simulium species No. 1 274 126. Collection location for Simulium species No. 1 in Florida. .275 127. Small flow to Pine Barrens Creek, Site 74, where Simulium speaies No. 1 was collected 276 128. Gametocytes (G) of L. smithi among normal turkey blood cells 291 129. Ookinetes of L. smithi 291 130. Sporozoites of L. smithi photographed in saline 292 131. Stained sporozoites of L. smithi 292

PAGE 13

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 BLACK FLIES (DIPTERA: SIMULIIDAE) OF FLORIDA AND THEIR INVOLVEMENT IN THE TRANSMISSION OF Leucocytozoon srrtithi TO TURKEYS 3y Dennis Drew Pinkovsky August, 1976 Chairman: Jerry F. Butler Major Department: Entomology and Nematology Immature and adult black flies (Diptera: Simuliidae) were collected in Florida over a period of three years. Eighteen species of black flies including ten which are new records for the State were found to occur in Florida. Records of black flies from 192 locations in 50 counties are included. Biological information is provided for each species together with data on distribution, seasonal occurrence, stream ecology and species associations. Keys to the Simuliidae of Florida are provided and structures are illustrated. Representative specimens have been deposited in the Florida State Collection of Arthropods and in the United States National Museum, Washington D.C. Transmission investigations have incriminated three species, Simuliwri congareenarum, S. meridionale, and 5. slossonae, as vectors of Leucocytozoon smiihi to turkeys in Florida. On nineteen occasions L. smithi was transmitted to domestic turkeys by the bites of infected black flies.

PAGE 14

CHAPTER I INTRODUCTION Members of the family Simuliidae as immatures inhabit flowing water and as adults are often blood feeders which may vector diseases to man and animals in many parts of the world. A knowledge of the composition, distribution, ecology, and habits of the simuliid fauna of any area of local concern is essential for proper assessment of the impact on man of these insects. Successful black fly control is dependent on thorough knowledge of the major breeding sites, seasonal occurrence and other facts concerning local Simuliidae. When I arrived in Florida to begin my Ph.D. research I wrote to the Florida Division of Health offices in Jacksonville, Vero Beach, and Panama City and enquired about previous investigations on the black flies of Florida. I found that little work had been done in the State on this family of biting flies (Beck, 1973; Linley, 1973; Rogers, 1973 all personal communications). Mrs. A.T. Slosson collected black flies in Florida (Dyar and Shannon, 1927), probably at the turn of the twentieth century. Some of Mrs. Slosson 's specimens and a portion of those of Calvin Jones, Harry Gouck, Darrell Anthony and other USDA researchers who collected black flies in the northern and central portions of the State in the 1940' s and 1950' s are located in the U.S. National Museum. Based at least in part on the examination of these specimens Stone (1965-) listed the following seven species from Florida: Cnephia (Cnephia) peeuarwn (Riley); Simulium (Eusimulium) -1-

PAGE 15

-2congareenarum(T)yar and Shannon); Simuliwn (Byssodon) meridionale Riley; Simulium (Byssodon) slossonae Dyar and Shannon; Simulium (Simuliwn) decorum Walker; Simulium (Simulium) jenningsi Malloch; and Simulium (Simulium) tuberosum (Lundstrom). Stone and Snoddy (1969) did not list S. jenningsi as occurring in Florida but did record two new species Simulium (Phosterodoros ) jonesi Stone and Snoddy and Simulium (Phosterodoros) nyssa Stone and Snoddy from the State. Thus the total number of species of black flies known to occur in Florida when I began my research was eight. This research was initiated in the fall of 1973 with the following objectives: 1) to determine the species complement, distribution and seasonal occurrence of black flies throughout Florida; 2) to gather ecological information primarily on the immature stages; 3) to provide keys with illustrations to the black flies of Florida; 4) to gather and deposit in the U.S. National Museum of Natural History and the Florida State Collection of Arthropods as complete a collection of adult and immature Florida black flies as possible; and 5) to determine the vectors of Leucocytozoon smithi in turkeys in Florida.

PAGE 16

CHAPTER II LITERATURE REVIEW Simuliidae Taxonomy and Distribution Rubtsov (1974) presents a concise discussion on the history and major advances in black fly taxonomy. Linnaeus (1758) first described two simuliids and assigned the names Culex reptans and Culex equinus but did not differentiate them from mosquitoes (Davies et al., 1962). Latreille created the name Simulium for the genus in 1802 using Rhagio oolombasohensis Fabricus as the type (Stone and Jamnback, 1955) . This remained the only genus for the thirty to forty black fly species described up to the turn of the twentieth century. The work of Smith and Kilbourne (1893) and others which focused on arthropods as vectors of diseases stimulated much research on insects and related groups of medical and veterinary importance. Roubaud (1906) designated two black fly subgenera, Prosimultum and Eusimulium, based on differences in wing venation. Lundstrom (1911) in Sweden and Jobbins-Pomeroy (1916) in the United States were early taxonomists who made use of the male terminalia of simuliids to separate species. Edwards (1915) in England also used male terminalia but more significantly in 1920 stressed the importance of studying black fly larvae and pupae. Dyar and Shannon (1927) first used female genitalia to separate North American black flies. Enderlein (1930) divided the family Simuliidae into 29 genera. Smart (1945) took a more conservative approach and distinguished 6 genera in the family: -3-

PAGE 17

-4Parasimulium Malloch, Prosimulium Roubaud, Austvosimulivjv. Tonnoir, Gigantodax Enderlein, Cnephia Edwards and Simuliwn Latreille. Stone (1963) listed 11 genera and 22 subgenera in the Simuliidae which he considered valid. Crosskey (1969) stressed a trinomial approach to simuliid taxonomy with heavy reliance on subgenera. Rubtsov (1974) recognized 17 genera of black flies in the Palaearctic region and listed 59 genera for the Simuliidae of the world, many of which American and British authors regard as subgenera. Brues et al. (1954) provide a worldwide bibliography of important papers which deal with the taxonomy of the Simuliidae up to the early 1950' s. In addition to the works already mentioned outstanding publications since about 1950 dealing with the classification and distribution of black flies outside of the United States include: Palaearctic region Davies (1968) Britain, Carlsson (1969) Spain, Rubtsov (1956 and 1962) U.S.S.R., Kuusela (1971) Finland, Zivkovic (1971) Yugoslavia, Rivosecchi (1971 and 1972) Italy, Crosskey and Peterson (1972), and Zwick (1974) Germany; Ethiopian region Crosskey (1957) West Africa, Travis et al. (1967), Crosskey (1969) Africa and its islands, Fain and Elsen (1973) Cameroons, Lewis and Raybould (1974) Tanzania; Oriental region — Travis and Labadan (1967a) Asia and European U.S.S.R., Delfinado (1969, 1971) Philippines, Crosskey (1973), Takaoka (1973) Nansei Is., Uemoto et al. (1973) Japan, Lewis (1973) Pakistan, Datta (1973, 1974, 1975) India; Australian Dumbleton (1963, 1972) Australia and New Zealand, Travis et al. (1968); Neotropical — Leon and Wygodzinsky (1953) Ecuador, Dalmat (1955) -

PAGE 18

-5Guatemala, Vulcano (1967), Travis and Labadan (1967b), Barreto (1969) Colombia, Wygodzinsky and Coscaron (1970, 1973), Wygodzinsky and Najera (1970), Wygodzinsky (1971) Northern Andes, Perez (1971) Venezuela, Rubtsov and Avila (1972) Cuba, Travis et al. (1974); Nearctic (excluding the U.S.) — Syme and Davies (1958), Davies et al. (1962) Ontario, Wood et al. (1963) Ontario, Travis et al. (1969), Peterson (1970) Canada and Alaska, Lewis and Bennett (1973) Newfoundland . In addition to the above works those of Rubtsov (1970, 1974) and Coscaron and Wygodzinsky (1973) mention the variability which has been observed in characters that are used for taxonomic determination of black flies. Gambarian and Terterian (1973) applied a numerical taxonomy approach and using 100 characters tried to separate simuliids in the Eusimuliwn group. The subgenus was found to be very homogeneous and creation of supraspecif ic taxa was not statistically justified. Internationally, black fly species determinations based on classical external morphological characters and behavioral differences are being supported by cytological, specifically chromosomal, studies. Rothfels (1956) provided an introduction and overview on analytical techniques for cytologically comparing black flies. Landau (1962), Dunbar (1969), and Vajime and Dunbar (1975) used chromosomal differences to separate forms or sibling species of a variety of simuliids including disease vectors where previously each had been considered a single species. Taxonomic works covering the whole United States include: Coquillett (1898, 1902), Malloch (1914), Jobbins-Pomeroy (1916), Dyar and Shannon (1927), and Stone (1965). Other publications were more regional in

PAGE 19

-6scope. Western and midwestern articles Include: Twlnn (1938), Smith and Lowe (1948) California; Nicholson and Mickel (1950) Minnesota; Stone (1952), Sommerman (1953) Alaska; Wirth and Stone (1956) California; Stone and DeFoliart (1959); Anderson and Dicke (1960) Wisconsin; Stone and Boreham (1965), Hall (1972, 1974) California; and Corredor (1975) Washington. Northeastern U.S. works include: Johannsen (1903), Leonard (1926), Metcalf (1932), DeFoliart (1951), Stone and Jamnback (1955), Jamnback and Stone (1957), Jamnback (1969), Pinkovsky (1970), and, reporting the unusual find of a South American black fly in the U.S. Wygodzinsky (1973) New York; O'Kane (1926) New Hampshire; Frost (1949) Pennsylvania; Dimond and Hart (1953) Rhode Is.; Sutherland and Darsie (1960a and b) Delaware; Stone (1964) Connecticut; Holbrook (1967) Massachusetts; Eckhart and Snetsinger (1969) Pennsylvania; and Crans and McCuiston (1970a) New Jersey. In the southeastern U.S. Tucker (1920) gave accounts of personal experiences with black flies and reported two species, S. pecuarum and S. meridionale , from Louisiana. Jones and Richey (1956) conducted a survey of the black flies of Jasper County, South Carolina, and discussed the biology, ecology and relationships to Leucocutozoon in turkeys of one Cnephia (pecuarum) and seven Simulivm species (congareenarum, decorum, jenningsi, slossonae, tuberosum, venustum and undescribed species). Snow et al. (1958) reported on the ecology, habits and distribution of black flies occurring in the Tennessee River Basin and recorded species in the three genera: Cnephia (mutata), Prosimulium (hirtipes, magnum, plus undescribed species,*, and Simulium (decorum, fibrin flatum , jenningsi 8U. group, meridionale, pictipes, tuberosum, venustum, verecundum.

PAGE 20

-7vittatum and three undescribed species J. Snoddy and Hays (1966) mentioned that eleven species of Simuliidae were captured at one location in Alabama using a New Jersey light trap modified for daylight use by removing the light and substituting, as an attractant, carbon dioxide gas dispensed at .45 kg (1 lb)/hr. Stone and Snoddy (1969) present distribution records, descriptions, and some biological information for 28 species of black flies discovered or expected to occur in Alabama. Garris et al. (1975) present the seasonal distribution of 7 species of Simulium collected from streams in Sumter Co. , South Carolina, and mention observations of black flies feeding on turkeys and captured in a C0 2 ~baited trap. Snoddy and Beshear (1968), and Snoddy (1971, 1976) describe and give some facts on the biology of three new species (S. taxodiwn, podostemi and lakei) in the expanding species list of the former S. jenningsi group. Bionomics The eggs of black flies are dropped freely into the water or are attached to substrates in the flows of streams and rivers. The larvae which develop feed by filtering material from the current or by scraping organic matter off the substrate and, usually within a month, transform into pupae. From the pupa which is normally attached to rocks or vegetation beneath the surface of the stream an adult fly emerges, in a bubble of gas, rises to the surface and is immediately capable of flight. The adult females of most black fly species suck blood and both male and female flies obtain energy from natural sugar sources such as nectar and honey dew. Eggs . Ussova (1961) describes most black fly eggs as between .24

PAGE 21

-8and .33 mm long and irregularly triangular in shape with rounded corners; the eggs of some Cnephia species, however, are said to be larger and elongate elliptical. Golini and Davies (1975) found black fly eggs were .228 mm long and .139 mm wide. Davies and Peterson (1956) examined the eggs of four genera (Gymnopais, Prosimulium, Cnephia, Simulium) and found no external sculpturing. The eggs are light in color when just laid and darken as the embryos develop. Davies and Peterson (1956) found that the eggs of Gymnopais and Prosimulium were the largest and those of Prosimulium and Cnephia were the narrowest of the genera checked. Black fly eggs are often found attached to vegetation and rocks in a swift flowing stream or river or lying free on the stream bottom. Tarshis (1968) conducted experiments on and reviewed accounts in the literature of desiccation and the overwintering of black fly eggs and concluded, as Wu (1930) had done earlier, that only those eggs which are kept moist, by damp stones or underground springs, etc., in apparently dry stream bottoms and other situations are able to remain viable. Tarshis (1968) found that freezing eggs of a number of species at to -70°C killed the embryos, while he was able to maintain moistened eggs alive for 424 days at 2-9°C. Kurtak (1974) found that eggs of Simulium pictipes , recovered while encased in ice from rock crevices along a stream, hatched in the lab after three days in 10°C flowing water. In northern areas eggs of many black fly species hatch when the water temperature reaches around 8°C (Carlsson, 1967) . Raybould and Grunewald (1975) found eggs of the Kibwezi form of S. damnosum hatch at 20°C four days after oviposition. Tarshis (1968) indicates normal egg hatch for Maryland black fly eggs occurs within one to five days. Larvae general morphology and deve lopment. Black fly larvae are usually between 4.5 and 10 mm long when mature, club-shaped with a wider

PAGE 22

-9poaterior end, and possess a pair of multiple-rayed cephalic fans on the anterior end, an unsegmented ventral proleg in the thoracic region, and a circlet of anal hooks posteriorly. Dark cephalic head spots mark the origins of cephalic muscles. Tarshis (1968) found that larvae hatched from eggs in the lab after 7-38 days at 10°C and 1-5 days at 20-25°C. Larvae developed to pupae in 18-50 days at 10°C and 11-29 days at 20-25°C. Cameron (1922), Dalmat (1955) and Reisen (1975) found six larval instars in the black flies they studied. Johnson and Pengelly (1970) observed S. rugglesi to pass through seven larval instars and Fredeen (1975) mentioned a seventh and final instar for S. araticwn. Craig (1975) distinguished nine larval instars in Tahitian species of black flies. Larval habitat . Larvae frequently attach by their posterior hooks onto a patch of silk secreted by their large, looped salivary glands. The silk is often applied to the same substrate on which the eggs are found. Larvae may drift a short distance tethered to the old substrate by a silk thread (Tarshis and Neil, 1970). A larva may also move in a looping, geometrid fashion from silk patch to silk patch alternately attaching its proleg then its anal disc (Dalmat, 1955). Elliot (1971) reported that black fly larvae can actively migrate upstream as well as passively downstream. Clean, smooth items free of algae or slime are preferred attachment substrates for the larvae and larvae are often located on such objects in stream sections where the current is increased by a partial obstruction (Dalmat, 1955; Carlsson, 1967). Larvae show positive phototaxis and colonize light substrates faster and more densely than dark substrates (Carlsson, 1967). Unusual attachment sites include the phoretic associations of black fly larvae on prawns, mayfly nymphs, crabs, and other arthopods reported by

PAGE 23

-10Disney (1971, 1973, 1975) and the attachment of young larvae of S. damnosum to older larvae of the same species (Burton, 1971). Carlsson (1967) in Sweden found the greatest concentrations of larvae in flows 80 to 120 cm /sec although some species preferred 40 cm/sec currents. Cariaso (1962) in the Philippines found immature black flies concentrated where the water velocity was .5-. 86 m/sec (1.63-2.83 ft/sec). Wu (1930) reported that simuliid larvae remained well established at a velocity of 1.83 m/sec (6 ft/sec). Rohdendorf (1974) suggested that the closed respiratory system of black fly larvae resulted after invasion of the swift flowing well oxygenated habitat typical of most simuliids. Tarshis (1968) found that black fly larvae could not live more than eight hours in still water. Anderson and Shemanchuk (1975), however, report they transported black fly larvae for several days in shallow, non-agitated, ice-chilled water with little mortality. Wu (1930) found approximately equal amounts of dissolved oxygen in quiet and turbulent sections of black fly streams and after a number of experiments concluded that black fly larvae have a definite requirement for current to lessen sedimentation and supply adequate food, not because of improved oxygen conditions. Crans and McCuiston (1970b) found larvae in permanent rivers and streams, temporary creeks and flowing roadside ditches. Dalmat (1955) and Lewis and Bennett (1975) reported finding a few live larvae in situations where the flow was so slow that silt and mud present prevented the larvae from anchoring to any fixed object. Van Someren (1944) reported S. ruficoime larvae from small pools and footprints in sandy river beds in Somaliland. Certain species are typically found in limited types of lotic habitats: S. aratiaum in large rivers (Cameron, 1922), S. piatipes at the outflow of dams and on

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-11flat bedrock (Snow et al., 1958) and S. oehraceum in very small streams about one meter (a few feet) wide and a few centimeters (a few inches) deep (Dalmat, 1955). Travis and Vargas (1970) found that in contrast to the clear mountain streams, the lower, slower Costa Rican streams most polluted with sewage and garbage had the greatest larval and pupal populations. Cariaso (1962) reported that black fly larvae were discouraged from breeding in water polluted with nitrogenous and human wastes. Carlsson (1967) indicates moderate pollution increases organic drift and is good for larvae but heavy pollution clogs the cephalic fans. Streams in Wisconsin carrying large amounts of eroded soil particles and other detritus were observed to be poor habitats for most black fly larvae due to feeding interference (Anderson and Dicke, 1960). Larvae feeding structures, behavior, and food . Davies (1974) and Craig (1974) describe the evolution and development of the important larval feeding structures in many species, the cephalic fans, and mention how these lateral palatal brushes which are well developed filtering devices in most black flies are apomorphically absent in later instars of Gyrnnopais and Twinnia and modified into raking structures in Cvozetia species. Chance (1970) stated that particles from about 1 to 350 microns in diameter, the majority in the 10 to 100 micron range, were ingested by four filter feeding species she studied. Larvae were found to attach within 10 centimeters of the water surface, extend their bodies into the flow and alternately open and close the fans combing particles from the fans when they were closed into the cibarium where a bolus was formed. Mulla and Lacey (1976) report larvae, with their heads pointed downstream, attach to silk deposits with their anal hooks, rotate the body 90° to 180° and open the cephalic fans to strain matter

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-12from the flow. Chance (1970) suggests that the horizontally operating mandibles, especially in non-filtering, grazing forms like Tuinnia, scrape organic food off the substrate. Rubtsov (1974) discusses the outer sclerite of the labium the submentum or hypostomum, an anteriorly serrate structure bearing setae which lies just cephalad of the gular notch and mentions that it is a tactile organ, is important in producing the larval silk strands, and that it may serve to scrape food from the substrate. In species which lack fans the submental teeth sometimes acquire a spatulate shape. Jobbins-Pomeroy (1916) reported Euglena and Spirogyra were important food items for black fly larvae but Cameron (1922) found diatoms formed the main components of the food. Anderson and Dicke (1960) found diatoms, other algae, and considerable inorganic material in the intestinal contents of black fly larvae. Carlsson (1967) listed as larval food bacteria, plankton, plant and animal parts, pollen and aerial fallout. Spring flooding leads to large outbreaks of black flies when rising water temperatures trigger synchronous hatching of many black fly eggs. The rising stream and river waters make more substrate available for larval attachment and increase organic debris and hence food available for the larvae. Bacteria filtered out by S. underhilli led to speculation of using simuliid larvae as indicators of pollution (Snoddy and Chipley, 1971). Reisen (1974) found that larvae remove bacteria and organic and inorganic particulate matter. Young larvae of Simuliwn species in nature fed at a faster rate than did older larvae as indicated by the time necessary to void a plug of dye particles (Mulla and Lacey, 1976). These authors also found that at a lower temperature (12.8°C = 55°F) a longer time (35-55 min) was necessary to

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-13eliminate a particulate plug from larval guts than at a higher temperature (30°C = 86°F, 20-30 min) . Pupae . Pupae attach to the same substrates as the larvae. The larvae form silken cocoons in which pupation occurs. Hinton (1958) mentions a pharate pupa stage where pupal characters are visible within the last larval skin but the larval mouth parts are still articulated and feeding continues up to when this "pupa" begins spinning the cocoon. The cocoons may be thin and rudimentary (Twinnia) , shapeless, irregular masses of silk (Pros intuition and some Cnephia) , sturdy, coarse cocoons (Simulium pictipes) or tapered, slipper-like finely shaped and tightly woven cocoons (most Simulium) (Stone and Jamnback, 1955). Members of the Simulium subgenus Phosterodoros have cocoons with forward edges that are convex in profile and have a large opening anteriorly on each side of the cocoon (Stone and Snoddy, 1969). Field and Low (1961) describe what Underhill (1944) illustrated for S. jenningsi (as S. nigroparvwn) , that is, sexual dimorphism in the cephalic plate of simuliid pupae. The cephalic plate, a sclerite that begins between the bases of the antennal sheaths and extends over the cephalic area, is longer and narrower in the male than it is in the female pupa. The pupae usually face downstream and are bordered on each side by tubular filaments which vary in number and shape according to species and aid in respiration. Cameron (1922) observed that the number of pupal filaments in S. avctiaum varied from the usual 12 to 13 or 11 and that one set might vary while the other group on the same pupa might bear the more typical number of filaments. Coscaron and Wygodzinsky (1973) mention the variation in the point of bifurcation or petiole lengths of the respiratory filaments of different specimens within the same species.

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-14Adults emergence . Tarshis (1968) reported that pupal development of three species of Simuliwn took 1 to 5 days at 20-25°C and that at 10°C It took 21 days following pupation before adults began emerging. Davies et al. (1962) describe how, at emergence, after gas fills the skin of the submerged pupa, the adult ruptures the pupal exuvium, pulls itself through a T-shaped opening and quickly floats to the surface in a bubble of gas (also see Hannay and Bond, 1971, section below). Carlsson (1967) found that a heavy silt cover on pupae prevented adults from emerging. Adults may easily be reared from pupae placed on moist filter paper in petri dishes (Hannay and Bond, 1971; Sutcliffe and Mclver, 1974). Wenk (1965) found males emerge sooner from pupae than do females. Disney (1969) found three species of Simulium in Africa pupate by day and may prolong pupation to avoid emerging at night. On warm days eclosion occurred early in the morning, on cooler days eclosion peaks occurred during the late morning and mid-day and under artificially cold days emergence was shifted to the late afternoon. Adults appearance and morphology . Adult black flies rarely exceed 5 mm in length (Dalmat, 1955). Male and female black flies are hump-backed in appearance. The males are holoptic, usually darker and more velvety than females and often bear a shiny pair of anterior lateral spots on the scutum (Davies et al., 1962). Costalization (i.e., a strengthening of the anterior wing veins and a decrease in wing venation in the posterior portion of the wing), shortening of the legs, evolution of the calcipala and pedisulcus, increase in head size and shortening of the abdomen with reduction of tergites and sternites are considered apomorphic developments (Rohdendorf, 1974; Rubtsov, 1974). Hungerford (1914) in discussing the anatomy of Simuliwn vittatum adults noted that

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-15black fly females possess a single spermatheca. Lewis (1957) presented morphological information about adults of S. damnosum and suggestions on dissection techniques. Bennett (1963b) found the shape and appearance of the salivary glands of adults valuable in differentiating species. Hannay and Bond (1971) found raised cylindrical buttons, each with a wax filament, between the macrotrichia on black fly wings and suggest these structures may aid the adult in keeping its wings dry during emergence. Sutcliffe and Mclver (1974) described cleaning hairs and combs of cuticular teeth on the metathoracic legs and cleaning hairs on the prothoracic legs which are used to clean the wings and head appendages, respectively. Adults attraction and trapping . Wirth and Stone (1956) indicate that male black flies are readily collected at light traps. Snow et al. (1958) captured adults at light, on vegetation, in cars and biting mammalian hosts. Fallis and Smith (1964) succeeded in using ether extracts of birds as attractants for simuliids. Anderson and DeFoliart (1961) used a variety of caged birds to attract ornithophilic black flies to traps in Wisconsin. Golini and Davies (1971) found that females of S. venustum fly upwind to a carbon dioxide source and cease flying upwind if the CO source is turned off. Snoddy and Hays (1966) and DeFoliart and Morris (1967) made use of the attractancy of C0 2 to capture simuliids. The silhouette of a target rather than the reflectance is more important in attracting black flies (Gillies, 1974). Peschken and Thorsteinson (1965) state that black flies are more attracted to stationary targets than to moving targets and simuliids show less discrimination between form and shape of three dimensional objects than some other hematophagous insects. Bradbury and Bennett (1974a)

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-16found bloodseeking black flies in the genera Prosimulium, Cnephia, and Simuliwn were more attracted to black, blue, and red than white or yellow, especially when the colors were of low visible reflectance. Thorsteinson et al. (1965) describe a tripod canopy-type Manitoba fly trap which uses a large black sphere target to make the trap visually attractive to bloodsucking diptera. Bradbury and Bennett (1974b) found that black flies could distinguish between targets based on their color, largely independent of carbon dioxide flow rates. Carlsson (1967) stated lakes, shoals, bogs and swamps act as "collecting mirrors" for female black flies seeking ovipositioning spots. Female black flies seeking oviposition sites oviposited more on green and yellow colored plastic strips in a stream than on red, purple, white or black strips (Golini and Davies, 1975). The Malaise trap, a final valuable collecting device for black flies and other insects, was conceived as a passive or random flight interception trap; however there is some evidence that the degree of color contrast between the trap and background vegetation is an attractance factor (Roberts, 1970 and 1972). Adult mating . Females of C. dacotensis have mature eggs at the end of the pupal stage and copulate with males on damp rocks soon after emergence. In general, however, simuliids can mate any time from emergence to oviposition time (Davies and Peterson, 1956). Davies and Peterson (1956) were unable to induce male black flies to mate by broadcasting the sound of the female wing beat frequency. Compared to males which do not form mating swarms, males with mating flights have larger eyes and enlarged upper facets. Snow et al. (1958) observed that males of S. piatipes flew upside down with their abdomens curved upward, contacted females as they flew by overhead, settled to a solid substrate,

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-17and copulated venter to venter with the female situated uppermost. The presence of about one male for every two hundred females of Cnephia mutata is considered evidence for parthenogenic reproduction (Davies and Peterson, 1956) . Downes (1965) reports Prosimulium .ursinwi is a maleless species in which the females do not even emerge from the pupa but disintegrate shedding the eggs into the stream. Adult feeding . Black flies have been collected from flowers (Davies and Peterson, 1956). Hocking (1953) stated that black flies and other northern biting flies obtain the energy for flight almost entirely from floral nectar. Lewis and Domoney (1966) reviewed the importance of sugar feeding on bloodsucking, autogeny, and parasite development and reported finding glucose, sucrose, fructose and other sugars in the crops of 101 wild caught Simulium. There are scattered reports of black flies being attracted to and feeding on cold-blooded animals (Hagen, 1883; Jobbins-Pomeroy, 1916; Smith, 1969), but of more importance are their ornithophilic, mammalophilic, and anthropophilic bloodsucking habits. Cnephia dacotensis 3 Gynmoipais holopticus, Twinnia tibblesi and a few other black fly species acquire sufficient nutrients during the larval stage to produce eggs and do not suck blood (Davies and Peterson, 1956; Shewell, 1957). Rohdendorf (1974) suggested that limited larval food stimulated the adult female black flies to hunt and feed on vertebrates. Cameron (1922) mentions the tenacity and persistence black flies exhibit when feeding and states that once the mouth parts are securely inserted into the skin the insect is not easily disturbed. Ussova (1961) states that the mandibles pierce a host's skin, the maxillae with recurved teeth anchor the proboscis in the skin, and alternating actions of the cibarial and pharyngeal pumps suck the blood into the esophagus.

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-18James and Harwood (1969) indicate simuliids are telmophages or pool feeders which lacerate blood vessels with the toothed, transversely operating mandibles and vertically operating maxillae. Davies and Peterson (1956) record a wide, natural host range for some species such as S. venustum duck, crow, heron, deer, and human. Others such as P. hirtipes (now three species) and S. deaovwn which feed on mammals in the field fed on birds when placed in vials on the avian species in the lab. Yang and Davies (1974) examined the salivary glands of three black flies and found an anticoagulant factor which keeps the blood fluid for movement into the gut. These authors also report an agglutin factor in flies at least twelve hours old. Adult egg development and egg laying . Cameron (1922) found development of the ovaries of female black flies followed a successful blood meal. Davies and Peterson (1956) describe females of several species with weak teeth or only hairs on the mandibles and maxillae. These flies do not suck blood and emerge with already mature eggs. Davies and Peterson (1956) indicate eggs develop five to twenty-one days after emergence with the longer time involving a prolonged blood meal search under natural conditions. Immature and mature eggs are found together in the ovaries which suggests at least two ovarian cycles (Cameron, 1922). Females may survive long enough for three batches of eggs but few probably live this long in the field (Davies and Peterson, 1956). Cameron (1922) reported that the eggs of Simulium simile (-S. avatiovm) are oviposited on rocks in large cake-like masses embedded in a soft gelatinous matrix. Davies and Peterson (1956) mention Prosimulium and Cnephia species which deposit eggs freely while flying down or across a stream. Stone and Jamnback (1955) indicate S. vittatum lays strings

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-19of eggs in a gelatinous matrix. Golini and Davies (1975) in Canada found that female black flies settled at the water line on trailing cattail leaves and deposited large (16 cm x 2 cm) irregular egg masses one to five layers deep in a gelatinous substance. Davies and Peterson (1956) found simuliids oviposited an average of 200 to 500 eggs per female while Golini and Davies (19 75) report an average of 417 eggs for each female. Adults life cycle, longevity, range, and general activity . Tarshis (1968) reports a 21 to 25 day period for mixed cultures of five Simulium species to develop into adults after eggs were placed in aquaria. Raybould and Grunewald (1975) found developmental time from egg to adult ranged from 18 to 50 days for African black flies. Dalmat (1955) in Guatemala used colored dyes and the releaserecapture technique and found marked female black flies survived as long as 85 days and traveled as far as 9.7 miles. Bennett (196.3a) tagged females of S. rugglesi by feeding them on ducks injected with a phosphorous-32 solution and recovered labelled flies up to 28 days later and 9.6 km distant following watercourse paths. West et al. (1968) exposed 32 black fly larvae and pupae to P-treated water and recovered radioactive flies as far as 33.5 km (20.8 mi) distant. Bennett and Fallis (1971) found S. euryadminioulum flew up to five miles from the release point and report the average life span of the females was at least two to three weeks. Hocking (1953) summarized published records of flight ranges which reached a maximum of 145 km for S. reptans colwnbaczense. Wellington (1974) reported frenzied activity in black flies which was apparently correlated with barometric pressure changes as a storm system approached. Cameron (1922) observed swarms of male black flies

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-20or males and females on warm, cloudy days with rain threatening or falling gently. Carlsson (1967) reports that rapid changes in temperature, air pressure, and light seem to increase the activity of all simuliid species but he also mentions even a light breeze reduced black fly activity considerably. Hocking (1953) found flight was continuous in Simuliwn species down as low as 12.8°C (55°F) . Concerning nocturnal behavior Dalmat (1955) observed that black flies in Guatemala move down to the base of plants as the sun sets and can be captured at night using a lantern and sheet when the insects emerge from their resting sites near the ground. Wolfe and Peterson (1960) describe climbing trees and sweep netting twenty-five feet off the ground to capture black flies at night in Quebec. At dawn Wolfe and Peterson observed simuliids flying down from the canopy. Recently in India researchers used light traps and found that black flies were active throughout the night with a peak of flight activity occurring around midnight (Datta and Dasgupta, 1974). Lab colonization . Black flies have been collected in nature in all stages and have been brought to the lab for experiments. Tarshis (1965a) describes techniques for collecting and shipping viable black fly eggs. Tarshis (1965b) and Tarshis and Adkins (1971) discuss collecting large numbers of black fly larvae on artificial cloth substrates and mention techniques for transporting larvae in aerated containers. Tarshis (1971) discusses rearing black fly pupae to adults in individual brass strainer cloth cylinders connected to aquarium air stone units. Tarshis (1972, 1973) describes field collection, laboratory rearing of immatures to adults, and successful feedings of females of C. ovnithophilia on ducklings in the lab.

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-21Wenk and Raybould (1972) point out that colonies of insects aid critical studies on their biology, provide adults free from infection for transmission work and can provide abundant material for investigations at times when natural populations are limited. Dunbar (1969) mentions that hybridization experiments supported by colonies of flies and sufficient knowledge of mating requirements can provide valuable information concerning the species status of insect forms. Dalmat (1955) found carbon dioxide stimulated feeding and oviposition and succeeded in inducing 40% of 65,000 S. oohmcewn, S. oallidwn, and S. metallicwn to feed and 20% to oviposit but no eggs developed into larvae. Field et al. (1967) found that females of S. vittatvm confined in a vial bit man, quail, and rabbits. By impaling a male on a minuten and brushing the male genitalia against the genitalia of the female coupling could be achieved. No female of S. vittatum which mated survived to oviposit and eggs deposited by ether, unmated, females in the artificial flows available failed to hatch. In the laboratory Wenk (1965) successfully reared, mated, blood fed and achieved oviposition of viable eggs with Boophthova erythroaephala. Wenk and Raybould (1972) working with the Kibwezi form of 5. damnosum likewise reared, mated, blood fed and obtained viable eggs from female black flies in the lab. Mating, which was also achieved with a member of the S. bovis complex, occurred in partially lighted emergence cages and was confirmed by the presence of sperm or spermatophores. Raybould and Grunewald (1975) review the literature on lab colonization of black flies and mention difficulties confronting researchers: inducing mating, inducing females to feed on blood sufficiently and consistently, erratic viability or hatchibility of eggs oviposited in the lab, and removing wastes from immature rearing setups.

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-22Al though a few black fly species have been Induced to complete every stage of their life cycle in the lab, they are still not considered to be successfully colonized. Damage Jamnback (1973) describes the black fly bite reaction: a hemorrhagic spot which develops into an itching wheal; sensitive individuals may suffer headache, fever, nausea, glandular enlargement, and bites around the eyes may cause swelling that results in obscured vision. Loomis et al. (1975) report black flies cause considerable irritation and tissue damage to horses' ears, head, neck, and belly, producing wounds and papules. S. arctioum, C. peauanm, and S. eolumbaczense are described as major pests of livestock (Jamnback, 1973). Fredeen (1974) mentions black fly outbreaks during 1944-1947 in Saskatchewan which resulted in the deaths of 1100 farm animals including cows, horses, hogs, and shorn sheep. Fredeen (1975) reports a S. arctiaum outbreak in 1972 along the Northern Saskatchewan River that killed at least 18 cattle. Black flies transmit Leucoaytozoon parasites to ducks (Fallis and Bennett, 1966), trypanosomes to ducks (Desser et al., 1975), and Leuooaytozoon parasites to turkeys (Skidmore, 1931; Noblet et al. , 1972) (see also the Leuooaytozoon smithi sections in the Literature Review and Results and Discussion below). Sudia et al. (1975) mention that epidemic VEE virus has been isolated from Simuliim species in Colombia and although biologic transmission has not been proven, mechanical transmission by contaminated mouth parts may be possible for at least 72 hours. Travis et al. (1974) report a finding of vesicular stomatitis virus in black flies in Colombia. Eastern encephalitis virus has been

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-23isolated from a pool of 100 unengorged S. meridionale which indicates this species may biologically transmit the disease (Anderson et al., 1961). Steelman (1976) reports monetary losses in cattle herds due to worm nodules caused by simuliid transmitted Onchocerca gutterosa amount to $500,000 each year in Australia. Onchocerca volvulus, the causative agent of blinding filariasis in man, is vectored by black flies and has been reported from Africa, Yemen, Mexico, Guatemala, Venezuela, Colombia, and Surinam; indigenous cases may also occur in Brazil (World Health Organization, 19 71; Travis et al., 1974; Raybould and Grunewald, 1975). Microfilaria of 0. volvulus previously were observed in the eyes and skin, however, Anderson et al. (1975a) found microfilaria in the urine, blood, and sputum of persons treated with diethylcarbamazine. Diurnal periodicity of 0. volvulus microfilaria in the skin has been shown to correspond with the peak feeding periods of important black fly vectors in Guatemala and Africa (Anderson et al., 1975b). Duke et al. (1975) found the transmission potentials of 0. volvulus along breeding rivers for S. damnosum in Africa so high that no communities could survive there. Control Physical control . Impoundage of rivers and the creation of reservoirs together with planned periodic cutoffs of discharge water has discouraged or eliminated black fly breeding along many stretches of rivers (Snow et al., 1958). Removing debris such as planks, tree limbs, trailing vegetation and other potential substrates for black fly larvae can help control simuliid population levels in streams (Jamnback, 1973).

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-24Chemical control . Carestia et al. (1974) reported that aerial applications of malathion, Dibrom , or Dibrom plus a repellent for adult black fly control were unsatisfactory. High winds, dropping temperatures and decreasing daylight interfered with adult fly activity and chemical effectiveness. Rapid reinfestation of adults from areas just outside the treated zone is another reason why most control efforts have been aimed at immature rather than adult black flies. Fairchild and Barreda (1945) observed the effectiveness of DDT as a black fly larvicide. Davis et al. (1957) discussed early attempts at evaluating the larvicidal properties of parathion dripped into streams, dieldrin applied by air, and DDT delivered by various ground techniques as well as by airplane. Evidence of the persistence of DDT in non-target organisms and decreased susceptibility of black fly larvae from streams with a history of DDT treatments stimulated a switch to less persistent materials (Fredeen et al., 1971; Jamnback and West, 1970). Travis et al. (1970) in field trough tests found Dibrom was outstanding in producing t mortality of S. pictipes larvae at .5 ppm and 1 ppm concentrations and R R stated Dibrom, Gardona and Ciodrin deserved more practical stream tests. 4 Jamnback and Frempong-Boadu (1966) found chemical formulations are most effective which give uniform distribution of the insecticide in the water and have a specific gravity of slightly less than 1.0 to keep the toxicant near the surface where most of the black fly larvae are located. Abate R (20%) in Panasol plus .5% Triton X-161 (.998 specific gravity) applied by aircraft eliminated larvae up to .8 km (.5 mi) downstream from the treatment point. Pelsue et al. (1970) chose Abate as a black fly larvicide due to its low mammalian, avian, and fish toxicity and achieved 100% control of larvae in three locations

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-25with .5 ppm delivered for 60 minutes. Detachment of larvae occurred within 24 hours and reinfestation was noted in 15 to 60 days. Kissam et al. (1975) reported 2% Abate Celatom granules delivered at 91 g AI/ .4 ha monthly reduced larval populations to zero and the populations only slowly built back up during the two weeks following each application. Fredeen (1974) mentions that methoxychlor, another black fly larvicide, is minimally toxic to vertebrates and methoxychlor and its metabolites are not concentrated in fish or other aquatic species. In Canada in 1969 a 15 minute injection of .2 ppm methoxychlor emulsifiable concentrate caused the disappearance of 96% of the S. avoticum larvae 32 km downstream. Fredeen (1975) reported that a 7.5 minute injection of methoxychlor at .6 ppm killed 100% of the older black fly instars 40 and 80 km below the treatment area. Younger black fly instars which are more susceptible were depleted by 96% at a site 161 km downstream from , the injection. Fredeen et al. (1975) indicate that methoxychlor adsorbs to suspended solids in the water and may act selectively against the filter-feeding black fly larvae. Following a methoxychlor application Wallace et al. (1973) noted increased drift but no eradication of nontarget organisms and Fredeen (1975) found that non-target organisms repopulated more densely than before the treatment. Investigations of the effect of insect developmental inhibitors on black flies indicate that significant reduction (75-100%) of adult emergence can be attained in the lab with at least three black fly species with Altosid at concentrations between .001 and 1 ppm (McKague and Wood, 1974; Dove and McKague, 1975). Fromraer et al. (1975) reported that in field evaluation tests in New York DEET-treated light mesh

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-26jackets were effective in reducing landings per five minutes from 404 on controls to 1.2 on personnel with treated jackets. Biological control . Cameron (1922) found that a fish of no food value called the common sucker fed on 5. simile (=S. aroticum) larvae and pupae. Peterson and Davies (1960) review the predators of black flies which include adult Empididae and Dolichopodidae, Tendipedidae larvae, Trichoptera, Formicidae, Odonata adults and naiads, and spiders. Sommerman (1962) reported empidid larvae fed on black fly larvae. Snoddy (1968) found the solitary wasp Oxybelus emarginatwn to be a predator of adult black flies. Peterson and Davies (1960) and Chutter (1972) report cannibalism among simuliid larvae. Carlsson (1967) indicates predators are unlikely to serve well as man-manipulated biological control agents though the tricopteran genera Hy dropsy che and Rhyaaophila might give good results. One of the earliest accounts of parasitism of black flies is that , of Strickland (1911) who reported a mermithid nematode that retarded the development of the larval histoblasts and a sporozoan (microsporidian) which caused distorted and swollen larvae. Jamnback (1973) states that eighteen microsporidia have been described from black flies. Spores are ingested by the larvae, sporoplasm invades suitable host cells which are often in the fat bodies, multiplication and the production of many spores occurs, and the host larvae commonly die. Black fly pupae and adults have been found infected and transovarian transmission is common. Jamnback (1973) also mentions a fungus Coelomyoidium simulii which occurs in black fly larvae and is usually fatal. Davies (1958) reports microsporidians, mites, and mermithid nematodes as parasites of Canadian black flies and mentions that mermithids

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-27are found in larvae, pupae and both sexes of the adults. From 15 to 60% of the females in emergence collections were found to be infected with mermithids and females in oviposition swarms were also found with mermithids. The infected females which attempt to oviposit may serve to disperse the mermithids and introduce them to additional stream environments. Welch (1964) indicated that the mermithid life cycle began with the consumption by black fly larvae of the mermithid, usually in the infective first juvenile stage. Molloy and Jamnback (1975) observed direct penetration of the black fly larval cuticle by preparasitic juveniles of Neomesomermis fluminalis , a mermithid which infects at least fourteen species of black flies in three genera. Exit of the parasite from the host is almost immediately fatal to the black fly. Anderson and DeFoliart (1962) report 49-93% parasitism by Isomermis and Gastromermis in one black fly species in Wisconsin. As a step toward the mass rearing of these biological control agents Bailey et al. (1974) discuss techniques for the mass collection of mermithid postparasites from field collected black fly larvae. Batson et al. (1976) reported an iridescent virus from black fly larvae in Wales but could associate no major fluctuations in the black fly population with the presence of the infected larvae. Florida Ecological Habitats Florida is a peninsula that lies between 80 and 88 degrees west longitude and in the same latitude belt as the Sahara, Arabian, and other large deserts. Exclusive of the Keys, the State extends roughly 644 km (400 mi) north and south along the peninsula and about the same distance east to west along the north coast of the Gulf of Mexico. Florida covers

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-28151,710 sq km (58,560 sq mi)'. The highest point in the State is 105 m (345 ft) above sea level, at Lakewood in Walton County. From Orlando south to Sebring another high section occurs and Iron Mountain near Lake Wales is located about 91.5 m (300 ft) above sea level (Morris, 1973). The earliest geological horizon is the Ocala limestone formed during the Eocene, 58 million years ago. Modern Florida made its first appearance during the Oligocene, 36 million years ago, as a small island about where the counties of Suwanee, Columbia, and Alachua are now located and then rose as a peninsula during the Miocene (Byers, 1930). There are three schemes which have been used to break Florida up into sections according to: 1) vegetation; 2) geology; and 3) edaphic factors. These schemes are presented to help illustrate the ecological diversity in Florida, which on the one hand provides a range of habitats for simuliid species and on the other hand restricts black fly breeding to certain more suitable regions of the state. Byers (1930) divided Florida into seven biotic areas based on dominant types of vegetation: a tropical hammock strip from St. Lucie County through Dade County along the east coast extending a few miles inland with cabbage palms, mahogany, ironwood, papaya, epiphytic bromeliads, plus other plants; a grass swamp area south and east of Lake Okeechobee; magnolia and temperate hammocks, primarily along the east coast from Flagler County to Indian River County and on the Gulf Coast from Wakulla through Hernando Counties — these occur on well drained but moisture holding soils with magnolia, holly, and red bay as dominant vegetation on the east coast and beech, elm, sweetgum, hickory, and oaks dominant in the west coast hammocks; a section of southeastern deciduous forest extending down into Leon, Liberty and Jefferson Counties; tree swamps in Collier, Columbia, Baker

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-29and Nassau Counties which include Big Cypress Swamp and other cypress and also black gum swamps; pine flatwoods along the Gulf west of Wakulla County, on the west coast of the peninsula from Pasco to Collier Counties, and in the Clay County and Putnam County region — these flatwoods occur on level, poorly drained land underlain by hard pan which results in an acid soil upon which grow palmetto, grass and pines; and, lastly, the southeast coniferous forest with long leaf, yellow, and slash pines and saw palmetto on well drained uplands. Cooke (1939) divided Florida into five physiographic areas: central highlands, Tallahassee hills, western highlands, Marianna lowlands and coastal lowlands. The central highlands, in the center of the peninsula from Baker County and Columbia County south to Lake Okeechobee, contain in their southern section thousands of lakes which in the north section are accompanied by numerous streams. The Tallahassee hills and western highlands are composed mainly of red clay with a relatively good number of streams. The coastal lowlands usually lie less than 30 m (100 ft) above sea level and include swampy areas such as the Florida Everglades and marshy areas along the east and west coasts of the state. The Marianna lowlands in Walton, Holmes, Washington and Jackson Counties contain few permanent flows and consist mainly of swamps, flatwoods, and sandy hills bearing pines. Davis (1967) and Smith et al. (1967) present six major land resource areas for Florida: a southern coastal plain; gulf coast flatwoods; central Florida ridge; Atlantic coast flatwoods; southern Florida flatwoods; and Everglades and associated areas. The southern coastal plain, occupying the northern half of the state from Escambia County to Madison County, is covered by forests of mixed hardwoods and pines on the lower

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-30areas and forests of long leaf pines and oaks on upland clay or well drained upland sand. The gulf coast f latwoods — a strip along the gulf side of the panhandle from Escambia County through Citrus County, the Atlantic coast flatwoods primarily in Columbia, Baker, Nassau and Bradford Counties in the north, and the south Florida flatwoods occupying most of the southern section of the peninsula outside of the Everglades consist of pine flatwoods and swamp forests (pines, palmetto, herbs, bay tree, laurel, gum, cypress) on poorly to very poorly drained and marshy soils. The central Florida ridge is located on well drained soil in the middle of the peninsula and includes forests of long leaf pine and xerophytic turkey oak (now mainly planted in citrus) and hardwood forests of mixed evergreen and deciduous hardwoods on rich upland soils. The Everglades are primarily sawgrass, Mariscus jamaicensis , on peat and muck soils. Average January temperatures range from 10-13°C (50-55°F) in the panhandle to 18-21°C (65-70°F) in the Everglades area. Average July temperatures fall between 27-29°C (80-84°F) throughout the state. Annual rainfall averages 134.62 cm (53 in) (Raisz, 1964). June through September or October is considered the rainy season in Florida (Byers, 1930) however Berner (1950) indicates that in the northwestern area rainfall is more evenly distributed throughout the year. There are seventeen first magnitude springs in Florida (Morris, 1973). Despite the heavy precipitation there are relatively few surface flows since much of the water moves in underground channels in the underlying limestone (Raisz, 1964). The vast majority of true streams and rivers are found north of Lake Okeechobee. Morris (1973) indicates there are 1,711 streams, rivers, and creeks in Florida with a total length of 16,989 km (10,550 mi).

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-31Berner (1950) states that in Florida there are relatively few intermittent streams. He divides permanently flowing streams in the State into: sand-bottom creeks with loose shifting sand and little vegetation; sand-bottom creeks with fairly loose sand and considerable vegetation; silt-bottomed creeks with little vegetation; and siltbottomed creeks with considerable vegetation. Stagnant rivers occur in south Florida where they serve primarily as drainage canals for the Everglades and though there is considerable vegetation along the shores of the rivers their fifteen to twenty-foot depths restrict plant growth toward the middle. Larger calcareous streams contain water which rises from springs, are basic in pH reaction, and in shallower sections are lined on the bottom with Vallisneria (eel grass), Sagittaria (arrowhead), algae and other plants. Erving (1971) notes that the temperature for Florida springs is about 22°C (72°F) year round. While noting springs are alkaline Berner (1950) also indicates that marsh and swamp water can be very acidic reaching a pH of 3.6 or below. He further adds slow flowing rivers to his classification of Florida's streams and rivers and included here are the Suwanee, Apalachicola and other rivers which have extensive drainage areas. Wood and Fernald (1974) list twentyeight major drainage basins and streams in Florida. The two largest, the Apalachicola River Basin and the Suwanee River Basin drain 50,777 sq km (19,600 sq mi) and 25,984 sq km (10,030 sq mi), respectively, some of which lies outside of Florida. Wood and Fernald (1974) indicate that peak annual flows occur in eastern rivers in September and October with buildups since June while more western rivers (Chipola, Apalachicola) reach maximum flow during March and April. In northwestern Florida a continental weather pattern predominates whereas a more

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-32tropical weather routine predominates in central and south Florida. Beck (1965) recognized five stream types in Florida: sand-bottomed, calcareous, swamp and bog, larger rivers, and canals. He points out that due to velocity, substrate, and spring discharges many of these types of flows can be recognized at different points along a single watercourse and cites the Suwanee River which is successively a swamp and bog stream, a sand-bottomed stream, a calcareous stream and a larger river of mixed origin. Beck (1965) classifies swift flow in Florida's streams as that velocity suitable for Plecoptera while simuliid populations indicate conditions of moderate velocity. Below these two levels the speed is considered low. A general progression is recognized with the swifter portions of upper rivers marked by eroding limestone and thick growths of moss, midpoints along rivers being sand-bottomed and lower, slower portions of rivers containing deposits of finer materials on the bottom. Beck (1965) lists the sand-bottomed stream as the most common type of stream in Florida. The fauna consists of Hydropsychid and Philopotamid caddisflies, simuliid larvae, Plecoptera, Stenonema mayflies, orthocladine chironomids, and Corydalis. In these sandbottomed streams the pH reaction is usually between 5.7 and 7.4 and they display moderate to high color and moderate to swift velocity. The bottoms consist of sand, leaf deposits, and limestone outcroppings, and plant growth may be dense. Beck mentions that swamp and bog streams are highly acidic (pH reaction 3.8-6.5), sluggish flows found in the coastal lowlands and central highlands with origins in swamps and marshes. Calcareous streams are cool, clear flows of spring origin with dense growths of vegetation. These flows are alkaline (7.0-8.2 in pH) , bear large mollusc populations, are generally of low color, of variable

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-33velocity and have sand, clay or limestone bottoms. The flows in the larger river category usually carry considerable silt and are turbid, have high and steep banks and bottoms of coarse sand and limestone. They demonstrate a pH reaction of 6.5-7.4 and have few shallow places and few aquatic plants. Beck (1965) mentions from about the area of the St. Lucie Canal to south of Homestead no natural streams of any consequence remain along the east coast of Florida and, instead, canals are prevalent although in the past the Miami River had a swift flow and rapids (Fairchild, 1976 — personal communication). The above discussion provides an introduction to the climatic, edaphic, biotic and hydrologic conditions which are found in the State of Florida. Leuaoeytozoon smithi During the 1880' s Danilewsky observed parasites which were later described and designated as species of Leuaoeytozoon and Haemoproteus (Bennett et al., 1965). Hsu et al. (1973) list 67 species of Leucooytozoon, one of which is reported from Meleagris gallopavo, the domestic and wild turkey. Theobald Smith in 1895 first reported a protozoan-parasite in the blood of turkeys in Massachusetts and Rhode Island (Smith, 1895). Similar gametocytes were observed in the blood of turkeys in France and named Haemamoeba smithi (Laveran and Lucet, 1905). The following taxonomic scheme which shows the position of the turkey parasite now known as Leucooytozoon smithi was modified from Aikawa and Sterling (1974) by crediting Sambon (1908) not Danilewsky (according to Bennett et al., 1975) with the genus Leuaoeytozoon'.

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-34Phylum — Protozoa Goldfuss, 1818, emend, von Siebold, 1845 Subphylum — Sporozoa Leuckart, 1879 Class — Telosporea, Schaudinn, 1900 Subclass — Coccidia Leuckart, 1879 Order — Eucoccida Leger and Duboscq, 1910 Suborder — Haemosporina Danilewsky, 1885 Family — Leucocytozooidae Fallis and Bennett, 1961 Genus — Leuaoaytozoon Sambon, 1908 Species — smithi (Laveran and Lucet, 1905). The following authors observed L. smithi in turkeys in their respective states or countries: Atchley (1951), Bierer (1954), Jones and Richey (1956) and Wehr (1962) in South Carolina; Savage and Isa (1945) and Bennett et al. (1965) in Canada; Volkmar (1929) in Minnesota and North Dakota; Hinshaw and McNeil (1943) in California; Johnson (1945) in Michigan; Kozicky (1948) in Pennsylvania; Skidmore (1931) in Nebraska; Stoddard et al. (1952) in Georgia; and Byrd (1959) in Virginia. Volkmar (1929) also lists L. smithi from Germany and the Crimean Peninsula and Cook (1971) lists Asia. Travis et al. (1939) reported Leuaoaytozoon smithi from wild and domestic turkeys in Florida as well as Georgia, Missouri, Alabama and South Carolina. Simpson et al. (1956) and Forrester et al. (1974) also reported Leuaoaytozoon smithi from Florida turkeys. Solis (1973) exposed curkeys, ringnecked pheasants, chickens, bobwhite quail, pekin ducks and chukar partridges in an area where L. smithi occurred and found that only turkeys were susceptible to L. smithi. Fallis et al. (1974) and Aikawa and Sterling (1974) present information on the life cycle and structure of the intracellular parasite, L. smithi. Leuaoaytozoon belongs to the same suborder, Haemosporina, as Plasmodium and Haemoproteus and undergoes a similar life cycle. Unlike Plasmodium, however, there is no erythrocytic schizogony and in addition to the red blood cells being invaded by merozoites as in Haemoproteus (and Plasmodium) , in Leuaoaytozoon white blood cells are

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-35also used as sites for gametogony (Huff, 1942; Fallis et al., 1974). Gametocytes, in the case of L. smithi, grow and split the host nucleus in two (Sambon, 1908; Volkmar, 1929). Plasmodium species are transmitted by mosquitoes and Haemoproteus (Farahaemopvoteus) species by the Hippoboscidae and Ceratopogonidae (Fallis and Bennett, 1961a and b; Bennett et al., 1974). Leucoeytozoon is known at present to be transmitted only by black flies. A parasite of chickens which is vectored by Culicoides was designated Akiba caulleryi by Bennett et al. (1965) but Akiba has recently been considered as a subgenus of Leucoeytozoon (Hsu et al., 1973; Fallis et al., 1974). Gametocytes are ingested with the blood of an infected turkey by a feeding black fly. Roller and Desser (1973) observed diurnal periodicity with L. simondi where peak gametocytemia corresponded with peak activity and biting periods of the vector. It has been suggested that there is periodicity of the gametocytes of L. smithi in the peripheral blood and correlation with the periodicity and feeding peaks of the vectors (Moore et al. , 1974). Early in the infection each gametocyte appears as a round body in the parasitized cells; later the gametocytes deform each blood cell into the characteristic spindle shape (Huff, 1942). Desser et al. (1970) found with L. simondi elongated cells resulted when merozoites developed in leucocytes and round cells resulted from invasion of red blood cells. Macrogametocytes stain more darkly with Glemsa stain than microgametocytes. Within minutes of ingestion the microgametocytes and macrogametocytes escape from their host blood cells. The microgametocytes exflagellate and syngaray occurs with the macrogametes. The zygotes formed grow into elongate ookinetes within twelve hours after the initial blood ingestion. Sections of flies have revealed ookinetes in the process of penetrating

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-36the midgut of the fly (Fallis et al., 1974). Fallis and Bennett (1961a) working with L. simondi detected only sluggish movement of ookinetes and expressed doubt that this motion could facilitate penetration of the gut wall of a black fly. Howard (1962) working with Plasmodium gallinaoewv observed no mobility or active penetration by zygotes. Howard suggests a largely passive incorporation into the gut wall to within 5 microns of the external basement membrane as the blood meal is digested and the distended gut returns from a squamous cell configuration to its original columnar cell morphology. Desser (1970) described a pore, strengthening struts, microtubules and elongate convoluted micronemes in the apical cap of the anterior end of L. simondi ookinetes and suggested these structures might aid penetration through the peritrophic membrane and into the fly midgut epithelium. In as little as 48 hours the ookinetes develop into oocysts containing 50-100 sporozoites just beneath the outer membrane of the gut. Sporozoites escape from the oocysts and penetrate and collect in the salivary glands. Desser (1970) describes structures on the anterior end of sporozoites of L. simondi which may contain proteolytic enzymes and aid in entering the salivary glands and, later, host liver cells. The sporozoites pass out the proboscis of the black fly with the salivary fluids when the fly bites another host. The sporozoites of L. smithi invade the liver parenchymal cells of the turkey and grow into hepatic schizonts. Huff (1942) reports with other species of Leuoooytozoon megaloschizonts in the heart, spleen and other host tissues. Such schizonts are not normally observed with L. smithi; however Siccardi et al . (1974) reported finding megaloschizonts in the kidneys of infected turkeys fed coccidiostat medication. The schizonts by asexual multiplication (schizogony) produce merozoites

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-37which enter red and white blood cells. Peters (1971) suggests that in the related genus Plasmodium merozoite penetration may involve proteolytic enzymes and be active or may be more passive with the merozoites being engulfed or invaginated into the blood cells. The merozoites grow into gametocytes in the blood cells and the cycle is completed. The prepatent period or time elapsed between entrance of the sporozoites and appearance of gametocytes in the peripheral blood is 10 to 16 days. Symptoms associated with leucocytozoonosis include: loss of appetite, emaciation, wheezing breathing, congested heart and lungs, drooling, drooping wings, sitting on the hocks, enlarged liver and spleen, anemia, impaired immunological responsiveness, and death (Wehr, 1962; Salsbury, 1971; Siccardi et al., 1974). Turkeys under 12 weeks of age are often severely affected but mortality in older birds has also been reported (Simpson et al. , 1956). Many birds with high parasitemias appear outwardly normal. Birds that do not die may continue to carry the parasite for as long as 1% years (Skidmore, 1931). Borg (1953), Simpson et al. (1956) and others suggest that the pathogenicity of L. smitht has not been proven conclusively and that L. smithi may be an additive debilitating factor which when combined with other factors such as blackhead, fowl cholera or leukosis results in bird mortality. Stoddard et al. (1952), Skidmore (1931), Savage and Isa (1945) and others however have observed flock mortality up to 75% where bacterial cultures and other tests revealed L. smithi as the sole disease organism present. Jones and Richey (1956) indicated annual death losses due to Leuoocytozoon in one county in South Carolina averaged 5%. In 1973,132 million turkeys were raised in the U.S. and farmers grossed $934 million (Poultry Digest, April 1974; Agricultural Situation,

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-38November 1974). In the absence of severe threats by disease this industry will continue to grow as the public's demand for poultry and its products like turkey rolls and t.v. dinners as an alternative to beef increases. A number of black flies have been incriminated as possible vectors of L. smithi. Skidmore (1931) incriminated S. ocaidentale ( =meridionale ) as a vector of L. smithi in Nebraska by grinding up a number of flies that had fed on infected turkeys, injecting them in saline into clean birds, and 12 days later observing gametocytes of L. smithi in the turkeys' blood streams. Johnson et al. (1938) in Virginia exposed uninfected turkeys to the bites of S. nigroparvum (=jenningsi) and obtained L. smithi transmission. Underhill (1944) further substantiated this work but Byrd (1959) was unable to infect clean turkeys by macerating and injecting S. jenningsi that had fed on diseased birds. Wehr (1953) by injection and Jones and Richey (1956) by fly-bite incriminated S. slossonae as a vector of L. smithi. Byrd (1959) succeeded in infecting turkeys with L. smithi by grinding and injecting females of Prosimuliwn hivtipes that had fed on infected birds. I have seen no further follow up of this work and Fallis et al. (1974) do not list any member of the former P. hivtipes complex as a vector of L. smithi. Noblet et al. (1972) ground up in saline and injected infected females of S. congareenarijm into turkeys and incriminated this species as a suitable or possible vector for L. smithi. Savage and Isa (1945) and Anthony and Richey (1958) have reported leucocytozoonosis in the supposed absence of simuliids although other biting flies such as Culicoides, Stomoxys, Diachlorus and mosquitoes were present. Control of leucocytozoonosis can be achieved by locating turkey flocks at least 12.8 km (8 mi) from known breeding locations of potential

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-39vectors and in areas free from possible wild turkey disease reservoirs. Control of black flies by large scale aerial treatment of streams with larvicides has been shown to markedly decrease the level of parasitemias of L. smithi in sentinel turkeys (Kissam et al. , 1975). Fine mesh wire screen enclosures might be a valuable preventive technique with smaller flocks. Clopidol, a coccidiostat , at .025 and .0125% in feed is effective in reducing the number of L. smithi parasites in turkeys as indicated by blood smears (Siccardi et al. , 1974). Investigators of leucccytozoonosis in turkeys in Florida have studied the presence, consequence, and distribution of L. smithi in the State but have not proven specific vectors. Travis (1939) reported that nearly 100% of the wild turkeys in low areas of Georgia and Florida were infected with L. smithi and suggested that an aquatic breeding insect might be involved. Simpson et al. (1956) investigating outbreaks of L. smithi in turkey flocks near Palatka, Florida, reported finding S. slossonae and a species #58 (=S. jonesi Stone and Snoddy, 1969) breeding near the turkey flocks and suggested that S. slossonae might be an important vector. Davis et al. (1957) reported S. slossonae as prevalent in Florida. Forrester et al. (1974) found a 72-75% incidence of L. smithi in 484 turkeys more than one month old sampled from 10 localities in Florida. These authors also noted that a drop in the infection rate of L. smithi observed in wild turkeys in the Sikes Fisheating Creek Wildlife Management Area and other areas of the State in 1971 corresponded with low rainfall and low stream conditions and theorized that reduced numbers of potential black fly vectors may have resulted in the reduced prevalence of Leuaocytozoon in turkeys.

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CHAPTER III MATERIALS AND METHODS Black Fly Survey Primarily, efforts in this survey concentrated on collecting larvae and pupae from Florida's streams and rivers. In Alachua County ten streams were selected and in nearby counties an additional twenty flows were chosen which were fairly permanent and represented a diversity of ecological conditions. An attempt was made to secure samples from each of these streams at least once a month. Approximately the same location was revisited each time to obtain some consistency in stream and substrate conditions in order to judge seasonal replacement phenomena and normal population changes. Other sites in south, southwest and west Florida were chosen, generally along highways or country roads for accessibility, which were visited at least quarterly to obtain records at all seasons of the year. At each location specimens were removed with forceps or placed still attached to small portions of grass, twigs and so on into four-dram lip vials with neoprene stoppers. Each vial contained 80% ethyl alcohol and a code number penciled on a small slip of paper. The iramatures, collected in 30 to 45 minutes were deemed sufficient to indicate numbers and species present. Dark pupae, indicating an advanced state of development, were placed in four-dram screw cap vials on a strip of paper toweling moistened in the stream and were held alive in order to allow the adults to emerge. Imagos could be reared very successfully in this manner from -40-

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-41older and frequently, even younger, more pale pupae. The forceps used were secured to my wrist by a string to prevent their loss when my balance was upset by the current, when I was wading and saw an occasional water moccasin, and at other critical times. A 1.53 m long rake and a modified, extendible, golf ball retriever were used to obtain substrate samples and specimens where it was impossible to wade. At each collection location stream dimensions, velocity, pH, temperature, color, and substrate were recorded on special data cards along with indications of the attachment substrates of the immatures and an estimate of the population size. Temperatures were obtained with a metal-cased pocket thermometer, range -1° to 49°C (30° to 120°F) . Initial attempts were made to determine the pH of the streams using two portable, electronic pH meters. After obtaining erratic, inconsistent results in the field with the meters, pH papers (pHydrion papers Micro Essential Labs, New York) were tried with more success. These pH papers come in ranges such as 3.5-5.5 and 6.0-8.0 between 1 and 12 pH units with illustrated color differences at .5 pH unit intervals and are rated accurate to .25 pH unit in buffered solutions. The papers were checked regularly for accuracy with pH 4.0, 6.5, 7.0 and 10.0 buffered solutions. Very acidic readings were confirmed by bringing chilled water samples into the lab and checking them on a Beckman pH meter. Experiments with electronic, propellered Gurley flow meters proved them too cumbersome to set up and difficult to use in the streams which were often shallow and clogged with submerged vegetation. The floating cork technique suggested by Dalmat (1955) and others was used. A weighted cork 3.8 cm in diameter which floated low in the water and was connected to a 3.05 m (10 ft) long string was timed with a stop watch to determine the stream velocity

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-42at different points where larvae and pupae were located. Most immature black flies were encountered on trailing vegetation at or near the surface and thus the floating cork velocity figures should reasonably reflect immature habitat conditions. A ruler one meter long with centimeter markings was used to determine stream depths and widths. The widths of larger flows were estimated by pacing across them, when possible, or by pacing across a nearby bridge. Other omnipresent items on collecting trips included an insect net and a snake bite kit. Alcohol specimens and pupal vials from each site were placed with the data card into cloth bags and transported to the lab where identifications were made with the aid of a Bausch and Lomb stereomicroscope. Larvae were usually identifiable without dissection (except unraveling of the respiratory histoblast); however representatives of each species were dissected and mandibles, respiratory histoblasts, cephalic apotome, gular notch, antennae and fans, and anal sclerite and hooks were prepared in cellosolve and mounted in balsam under individual small cover slips. Adults which emerged were allowed to harden for a few hours and were then killed and placed in alcohol, prepared and mounted in balsam on slides, or pinned on minutin nadeln inserted in white polyporous pieces on #3 insect pins. In addition to immatures, some adult collecting was undertaken using a variety of traps. Black fly adults have been obtained in Florida by other individuals using light traps and Malaise (flight) traps. In this research three types of adult fly traps were used: the Manitoba trap (Thorsteinson et al., 1965), the blackout box trap (Anderson and Dicke, I960), and a modified ramp-type trap. The latter two types of traps will be discussed with the Leucocytozoon work. A modified Manitoba

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-43trap (Fig. 1) was constructed using 3 bamboo poles, each 2.1 m (7 ft) long, with a hole drilled vertically into the uppermost node of each pole into which a leg of a metal tripod laboratory ring stand was inserted. A triangular serex or organza canopy 1.2 m (4 ft) tall was positioned inside the leg frame and anchored at the three lower corners by strings tied through a hole in each pole. The apex of the tapering canopy with a 5.1 cm (2 in) wide elastic opening was drawn up a short distance through the ring opening and stretched around the lower lip of a 17.8 cm (7 in) tall lantern jar, the collecting vessel, which rested on the ring. The jar was topped with a double layer of white serex or organza held in place with rubber bands. The jar contained an inverted plastic funnel affixed to the jar with hot plastic glue. On strings attached around the base of the jar and hanging into the canopy was suspended a 1.2 m (4 ft) long cord with a black, cylindrical (20 cm wide, 22 cm tall) metal can or a black cardboard triangle, 63 cm on each side, on the end. A 23 cm wide black plastic "skirt" was sometimes fitted over the top of the canopy and positioned at the lower margin of the canopy to accentuate the "window" effect. A 31 cm long cloth bag which contained 1.36-1.8 kg of dry ice was also hung inside the canopy. The dry ice and the black target attracted Simuliidae and other Diptera which flew or walked up the canopy into the collecting jar. Adults could be aspirated from the jar by removing the outer cloth layer and inserting an aspirator tube through a small slit in the lower layer of material. Specimens could also be taken to the lab in the glass jar and immobilized by cold in a refrigerator, though condensation inside the jar often wet the specimens. Manitoba trapping was accomplished using one or two traps at a time in six counties at twelve locations for one

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-44Figure 1. A modified Manitoba trap with a black plastic skirt.

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-45to eight hour periods during the months of January through October. The air temperature, wind speed and relative humidity were measured during the collecting periods. Trapped adults were pinned or mounted on slides as with reared adults. Identification of immatures and adults was possible using taxonomic keys in Stone and Snoddy (1969), Stone (1964) and smaller publications on individual species. Leucocytozcon smithi Transmission In the investigation of the transmission of L. smithi to turkeys in Florida, modifications of Koch's postulates were used as guidelines. The first aim was to establish an association in nature between the host, the disease, and the vector. It was necessary to have a suspected vector, preferably a clean, reared one, feed on an infected host in the lab and to observe the stages of the parasite, especially the infective stage, the sporozoite, in the fly. A clean host was then to ' be infected by a vector which had become infected in the lab with the Leuaooytozoon parasite being recovered in the gametocyte stage from the formerly uninfected turkey. In order to confirm the reported presence of L. smithi in certain areas such as the Lochloosa Creek and Fisheating Creek Wildlife Management areas (L. Williams, 1974 — personal communication; Forrester et al., 1974), I set out sentinel turkeys. The same was done to establish the presence of the disease in other areas, and to obtain infected hosts to serve as donors for transmission studies. These turkeys, like all clean poults used in the transmission studies, were either hatched from fertile eggs in a fly-proof room or obtained as disease-free one-day old poults and held in a fly-proof room until use. Difficulties in obtaining clean

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-46poults for experiments early in 1975 when winter and early spring species, suspected to be L. smithi vectors, were flying were overcome for work in early 1976. Usually three birds were placed out in the field near a black fly stream in a cage for about a week and provided with food and water (Fig. 2). There were three primary sentinel sites, all in Alachua Co., and additional birds were also set out for short periods at Fisheating Creek, Glades Co. (see Table 5). Sentinels were also placed out on a few occasions in a ramp trap described below. In addition I worked with Dr. D.F. Forrester at Fisheating Creek during 1974 when he exposed many young turkeys for two week periods in cages in cypress trees and on the ground and in 1975 when cages and ramp traps were used for turkey exposures. To determine the species of black flies generally present at the sentinel sites, immature stages were collected and adults on the wing were sampled with Manitoba traps as described in the previous section. To identify black flies specifically attracted to turkeys I used blackout box traps (Fig. 3), ramp traps (Fig. 4) and exposed turkeys (Fig. 5). Anderson and DeFoliart (1961) used the technique of blackout box trapping in Wisconsin. In Florida the blackout box traps were used at two locations in 1974 and an additional seven sites during 1975 (see Table 7). A turkey was exposed in a cage on the ground or on a platform in a tree for about 15 minutes. A large cardboard box with two plastic collecting cups on the top which appeared as bright areas inside the otherwise dark box was then placed over the cage and bird. Black flies completing their blood meals or unfed but on the turkey exhibited a positive phototactic response, flew to the collecting cups and were captured. The flies were removed to holding cartons, the turkey was re-exposed, and the process was repeated.

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-47— ' > * mm mn^ far****J^p £***>. Figure 2. Sentinel turkeys in an exposure cage. Figure 3. A blackout box trap in the field.

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-48tJPMUL t *-^u&j&ft Figure 4. One view of a ramp trap. *** . * "V

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-49An omnidirectional ramp trap was used with the hopes that black flies would be captured without the constant presence of the collector. The trap was used at three locations from April to July 1975 (see Table 6). The ramp trap was a wooden-framed cube approximately .92 m (3 ft) in each demension with 4 slanted lower level ingress panels of aluminum framing and organza material and 4 upper, vertical organza-paneled windows. A cage with host animals was placed and suspended inside the trap through a hinged access opening cut into the solid plywood top. The attracted insects would fly or bounce up the slanted ingress openings, enter the trap to feed on the hosts, theoretically be unable to escape and later be aspirated from the inside of the bright vertical cloth panels or windows. By capturing flies off normally one or two exposed turkeys it was possible to tell which species approached the birds and, more significantly, to observe which species fed on the birds. Capturing flies off exposed turkeys first involved immobilizing the host. Masking tape was wrapped around the turkey's legs just above the feet which were thus held together. The bird was placed on its side and a layer of gauze followed by a strip of wide masking tape was placed across the turkey's neck, chest, and legs and affixed to the platform which was usually a large plastic tray. The upper wing was propped up to provide the black flies with easy access to bare skin. Using this technique it was possible to monitor the number and type of ornithophilic adults present and capture them as well as observe their feeding locations, duration and behavior. Their feeding could also be assisted and securing of blood-fed flies from exposed infected turkeys could be more readily ensured. To do this, flies that landed on the feathers of the wing or

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-50elsewhere were quickly aspirated before they could scurry beneath the feathers and were placed in the glass aspirator tube against the bare skin of the underside of the wing elbow, the bare skin of the chest over the rib cage or against the turkey's neck and allowed to feed to repletion. The fly was observed, the feeding timed, and the turkey soothed and kept as still as possible to assure a complete blood meal. Once fed, the flies were transfered to paper pint ice cream cartons with netting tops, provided a moistened paper towel to maintain high humidity and transported to the lab. The exposed turkey technique was used to collect black flies on numerous occasions from January through August at 8 locations in 3 counties (Table 8). To obtain black flies that had not been exposed to infections in the field and to obtain specimens of known identity, especially when working with species in the Simuliwn subgenus Phosterodoros which are difficult to distinguish as adults and are much easier to separate as pupae, adults were reared in the lab. Some adults were reared in pupal vials from field collected pupae and other adults were reared form field collected larvae and pupae in an aquarium with aerated water and a fine mesh netting cover. Black flies which fed on exposed infected hosts in the field were held in the lab for at least two days after feeding and then allowed to bite an uninfected bird. All adult flies maintained in the lab were held in paper pint-sized ice cream cartons covered with netting and provided with moist cotton for water and a raisin for nutrition. Up to four cartons were kept at one time together in either a large plasticcovered aquarium or a glass desiccator-type container (Fig. 6) where high humidity was maintained by including a sponge half immersed in a

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-51Figure 6. Glass container and paper cartons used for holding black fly adults alive in the laboratory.

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-52cup of water. Flies captured in an unfed condition in the field or reared in the lab were allowed to feed on an infected turkey, held for two to seven days and then allowed to feed on a clean bird. All laboratory feedings involving uninfected hosts were conducted in a flyproofed former bull room at the Veterinary Science area at the University of Florida. This room was incandescent lighted, had a double door entrance and had windows covered with very fine mesh organza material. Feedings on infected hosts were conducted either in the Veterinary Science room or at the fluorescent-lighted Veterinary Entomology Lab on campus. Black flies were routinely taken off their sugar source at least 24 hours before planned feeding trials and maintained only on the water provided by the moist cotton. Black flies were aspirated from the cartons or chilled in a refrigerator and removed with forceps and placed, usually one per vial, into seven-dram, clear plastic, cylindrical vials covered with green, fine mesh (30/6.45 sq cm) netting at both ends and held against the bare skin of the immobilized turkey. Room temperature, relative humidity and feeding times were recorded. Uninfected turkeys bitten by possible vectors were held in a flyproof room and the appearance of gametocytes in the peripheral blood was monitored by blood smears obtained by wing vein punctures every 3 to 4 days. Blood smears were processed by air drying, fixing in absolute methanol and staining in a 1:50 dilution of Giemsa solution and distilled water. Flies were dissected in .9% physiological saline and midguts and salivary glands were observed for the presence of oocysts and sporozoites of L. smithi. A few midguts were fixed in 10% formalin, stained in Haematoxylin and mounted in balsam, with the necessary intermediate steps for dehydration and clearing, in attempts

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-53to more readily view the oocysts. Most midguts were air dried, fixed in methanol and stained with Giemsa, as were the salivary glands. The preparations were covered with permount and a cover slip for microscopic examination. Other fly structures were placed in 10% NaOH (except the wings) for 48 hours, rinsed in water and acid alcohol, placed' in 100% cellosolve for 15 to 30 minutes and mounted in balsam on a microscope slide.

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CHAPTER IV RESULTS AND DISCUSSION Simuliidae General Comments The primary efforts of this survey and ecological investigation of the black flies of Florida were concentrated on the immature stages. Over 700 adults were reared and pinned or mounted in balsam on slides and a number of adults have been captured in traps or on host animals (Tables 4, 6, 7 and 8) however the vast majority of specimens are alcohol -preserved larvae and pupae. Over 1,100 four-dram vials containing about 50,000 specimens from 1,080 individual, positive collections which have been examined and identified are on hand. In addition, records were obtained for Florida black flies from institution and individual collections and are included under the appropriate species. Table 1 is a list of the Simuliidae collected during this research in Florida. Eighteen species representing two genera and six subgenera are recorded. Figure 7 shows the 192 locations where I personally collected specimens or obtained records from the collections of other individuals. One star normally designates a single collection site; in a few cases a single star is used to indicate two collection sites adjacent to each other such as a main stream and a side drainage flow to that main stream. Each collection location is listed by county and site number in the Appendix and the name and location of each stream is given. The 50 counties from which black flies are recorded are listed in Table 2 -54-

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-55Table 1. Florida black fly species. Cnephia (Cnephia) ornithophilia Davies, Peterson, and Wood* Simulium (Byssodon) meridionale Riley Simulium (Byssodon) slossonae Dyar and Shannon Simulium (Eusimulium) congareenarum (Dyar and Shannon) Simulium (Phosterodoros ) dixiense Stone and Snoddy* Simulium (Phosterodoros) haysi Stone and Snoddy* Simulium (Phosterodoros) jenningsi Malloch Simulium (Phosterodoros) jonesi Stone and Snoddy Simulium (Phosterodoros) lakei Snoddy* Simulium (Phosterodoros) notiale Stone and Snoddy* Simulium (Phosterodoros) nyssa Stone and Snoddy Simulium (Phosterodoros ) taxodium Snoddy and Beshear* Simulium (Psilozia) vittatum Zetterstedt* Simulium (Simulium) decorum Walker Simulium (Simulium) tuberosum (Lundstrom) Simulium (Simulium) vereoundum Stone and Jamnback* Cnephia species undetermined No. 1* Simulium species undetermined No. 1* * New collection record for Florida.

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-56rJ J4&Figure 7. Locations in Florida where black flies have been collected.

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-57Table 2. Florida black fly distribution records by county.

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-58Table 2. (Continued) Species « g S a to ca ?h CO ES 3 r <3'0'0 ooSc^to cjj ca ssgwscc ^carOOV^b^oaQ^ti'^CaSCaCufi, County

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-59Table 3. Black fly associations based on collections of immature stages.* T3 V n a £ ^ -* S o> k *<* to <3i M K K-^a"T3+^^ to 32 26 14 4 2 2 32 10 1 32 84 3 1 38 107 73 52 15 186 55 26 84 2 19 7 6 2 49 23 3 3 24 21 4 3 5 5 1 38 2 19 92 73 4 21 8 4 1 107 19 24 5 19 36 2 1 40 11 3 253 44 4 73 7 92 36 150 15 54 19 2 2 3 1 1 1 2 52 6 73 40 150 8 44 10 2 15 2 4 11 15 8 9 50 33 3 2 9 6 7 32 186 49 21 5 21 253 54 3 1 44 50 6 78 1 10 55 23 4 8 44 19 1 10 33 7 78 3. 1 1 c.

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-60Table 4. Black flies captured in Manitoba traps. Date Location Collection times Species (all females) 1974: 3 May 30 May 28 June 28 June 6 July 11-12 July 16 July 16 July 19 July 19 July 24 July 26-27 July 15 Aug 17 Aug 15 Sep 1975: 24 Jan Univ. of Florida, Site 34 Hatchet Creek Preserve, Site 24 Lochloosa Creek, Site 22 Hatchet Creek Preserve, Site 24 Junction SR 225/ 340, Site 8 Fisheating Creek, Site 95 Lochloosa Creek, Site 21 Hatchet Creek Preserve, Site 24 Sandy Hatchet Creek, Site 1 Univ. of Florida, Site 34 Hatchet Creek Preserve, Site 24 Fisheating Creek, Site 95 Lochloosa Creek, Site 22 Univ. of Florida, Site 34 Lochloosa Creek, Site 22 Lochloosa Creek, Site 22 1730-2030 S. vittatwn 1130-1930 S. slossonae, S. (Phosterodoros) spp. 1800-2000 S. slossonae 1710-2020 S. slossonae 1500-1745 S. slossonae 0630-0930; 3. slossonae, 1730-2045 S. (Phosterodoros) spp. 1630-1845 S. slossonae, S. (Phosterodoros) spp. 1600-1800 3. slossonae, S. (Phosterodoros) spp. 1600-1730 S. slossonae 1500-1830 3. slossonae 1530-1815 S. slossonae 0700-1010; S. slossonae, 1530-1800 S. (Phosterodoros) spp. 1730-1845 S. slossonae, 3. (Phosterodoros) spp. 0830-1430 3. slossonae 1630-1805 S. slossonae, S. (Phosterodoros) spp. 1445-1710 S. eongareettarun, S. slossonae, S. (Phosterodoros) spp.

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-61Table 4. (Continued) Date

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Table 4. (Continued) -62Date Location Collection times Species (all females) 1976: 24 Mar Turkey Creek, Site 216 1640-1830 S. congareenarum, S. slossonae 26 Mar Sante Fe College, 1030-1800 S. tuberosum, Site 28 S. slossonae

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-63C. (C.) ornithophilia S. (P.) hay si 1 5. (B. ) meridionale i i S. (B.) slossonae S. (E. ) congareenantm S. (P. ) dixiense S. (P. ) jenningsi S. (P. ) jonesi S. (P.) lakei S. (P. ) notiale S. (P. ) nyssa S. (P. ) taxodium S. (P. ) vittatum S. (.S.) decorum S. (S. ) tuberosum S. (S. ) verecundum C. spp. No. 1 S. spp. No. 1 S" N D J F Months Figure 8. Seasonal occurrence of black flies in Florida.

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-64with an indication of the species found in each county. Table 3 presents information on the frequency of association of black flies in Florida's streams and rivers. Figure 8 shows the seasonal occurrence patterns of the collected Simuliidae. Table 4 presents information on species captured in Manitoba traps. Further reference to these figures and tables will be made in discussing the individual species below. Keys to the black flies of Florida, in all stages, are included. Introduction to the Black Fly Keys Characters which are important taxonomically on black fly larvae and are noted on Figures 9, 10, and 11 include: the anterior, multiple-rayed cephalic fans and their stalks (CF,CFS); the antennae (A); the central dorsal section of the larval head capsule referred to as the front oclypeus or the cephalic apotome (CA) which bears usually dark pigmented areas called head spots (HS) ; the ventral, setae bearing and anteriorly toothed hypostomium or submentum (S); posterior to the submentum is the throat (or postgenal) cleft (TC) or gular notch which differs in size and shape in the different species; laterally on the thoracic region are a pair of organs called the respiratory histoblasts (RH) consisting of a number of coiled filaments whose number and branching pattern are often diagnostic; the ventral proleg (VP); dorsally at the posterior end the anal gills (AG), osmoregulatory organs, protrude and may be simple and branchless stalks or may have many branches and be arborescent; behind the anal gills is located the anal sclerite (AS) or X-piece which assists the larva in releasing its posterior from the silk attachment patches; the circlet or many rows of tiny anal hooks (AH) at

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-65CF .V CFS . CA Figure 9. Dorsum of the head capsule of a black fly larva (S. slossonae, "Ve< ' ', Figure 10. Venter of the head capsule of a black fly larva (C. ormitkophi Via) .

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-66Figure 11. Lateral view of two black fly larvae (S. dixiense) Figure 12. Black fly pupa and cocoon (S. dixiense) ,

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-67the very end of the larva; and the ventral protrusions which occur on seme larvae and are called ventral or anal tubercles (VT) . Characters important on the pupa include: the filamented repiratory organs (RO) ; tiny setae-like structures on the thorax called trichomes ', and posterior dorsal tail hooks (Fig. 12). On the cocoon lateral apertures (LA) or windows occur anteriorly in the Phosterodoros species and the texture and general regularity of the cocoon is sometimes useful in separating species. On the adult wing the presence or absence of hairs under the subcostal vein (SC); hairs on the dorsal, basal portion of the radius (R) ; the presence or absence of a basal cell (BC) ; and color of hairs on the stem vein (SV) are valuable characters (Fig. 13). On the adult female head the shinlness or pollinosity of the frons (F) and its shape and that of the clypeus (C) , the color of the antennal segments (A) , and the shape or size of the sensory vescicle of the maxillary palps (SV) are useful taxonomically (Fig. 14). Important characters of the male genitalia include: the appearance of the ventral plate (VP) and the shape of its median portion; the shape and relative lengths of the basimere (B) and distimere or clasper (D) ; the presence or absence of a basal lobe on the distimere; and the number of distal spines (DS) or teeth on the distimere (Fig. 15). On the female hind leg the presence or absence of a dorsal groove called the pedisulcus (P) on the second tarsal segment and the size of the calcipala (CL), a flattened lobe on the inner side and at the apex of the basitarsal segment, as well as presence or absence of a basal tooth (BT) on the tarsal claws are important characters (Fig. 16). Structures useful for determinations in the region of the female

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-68Figure 13. A wing of the black fly Cnephia ornithophilia. Figure 14. A f rental view of the head of a female black fly (5, nctidle)

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-69Figure 15. The male genitalia of a black fly, Cnephia ornithcphilia. Figure 16. The distal portion cf the hind leg of a S. meridional e female.

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-70l\ Figure 17. The terminalia of a female S. meridionale.

PAGE 84

-71genitalia (Fig. 17) are the size and shape of the genital fork stem (GFS) and arms (GFA) , the ovipositor lobes, each anal lobe (AL) and each cercus (CR) . AJke v t o the larvae of th e_ black flics o f Florida . (Photographic illustrations of certain structures referred to in this key and the keys which follow are included in the sections on the individual species. The larval key is primarily of value for later instars. For further illustrations refer to Stone and Jamnback (1955), Davies et al. (1962), Wood et al. (1.963), Stone (1964), Stone and Snoddy (1969), Snoddy and Beshear (1968) and Snoddy (1971, 1976). la. Hypostoraium convex along anterior margin; head spot pattern as in Fig. 18 C. cjviithophilia lb. Hypostonrium concave or level along the anterior margin; the head spot pattern not as in Fig. 18 2 2a. (lb) Head spots light (white) 3 2b. bead spots dark or indistinct in a fulvous pattern 5 3a. (2a) Head spots consisting of a central, posterior white spot with dark rays projecting and diverging anteriorly (Fig. 247, Stone and Snoddy, 1969; no mature larvae collected), S. meridionals 3b. Head spots with anterior and posterior medial and lateral spots all white, spots visible in the usual positions 4 4a. (3b) Medial anterior and posterior white head spots bordered by a dark fulvous area on each side (Fig. 99); respiratory histoblast with 8 filaments 5. decorum 4b. No distinct dark fulvous border by central head spots; histoblast with 6 filaments S. vcvccundivn

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-725a. (2b) Gular notch in the form of a shallow inverted -v; headspots as in Fig. 121 C. species No. 1 5b. Gular notch rectangular, sub-rectangular or sagittiform 6 6a. (5b) Gular notch broadly sagittiform, extending at least across H of the venter of the head capsule, as wide as long 7 6b. Gular notch not sagittiform or else longer than wide 13 7a. (6a) Respiracory histob.lasts consisting of 10 filaments 9 of which th arise from a thick basal 10 ' (Fig. 60); anterior medial head spots often weak S. jonesi 7b. Respiratory histoblast not as above; all head spots usually dark, distinct 8 8a. (7b) Respiratory histoblast with 6 filaments; anal tubercles not prominent S. notiale 8b. Respiratory histoblast with more than 6 filaments; anal tubercles prominent • 9 9a. (8b) Respiratory histoblast with 7 filaments S. haysi 9b. Respiratory histoblast with more than 7 filaments 10 10a. (9b) Respiratory histoblast with 8 filaments S. taxodiwn 10b. Respiratory histoblast with more than 8 filaments 11 11a. (10b) Respiratory histoblast with 9 filaments S. lakei lib. Respirator} histoblast with 10 filaments 12 12a. (lib) 10 respiratory histoblast filaments with a pattern as in Fig. 81; anal tubercles prominent and conical S. nyssa 12b. 3 respiratory histoblast filaments with a pattern as in Fig. 53; anal tubercles small and rounded 5. jenningsi

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-7313a. (6b) Gular notch long extending % the distance or more to the teeth of the submentum, sagittiform or subrectangular , pointed or rounded anteriorly 14 13b. Gular notch extending less than h the distance to the teeth of submentum, subrectangular 16 14a. (13a) Head spots indistinct in fulvous area; gular notch usually with parallel sides; anal tubercles inconspicuous . . S. tuberosum 14b. Head spots distinct; gular notch with curved margins, elongate sagittiform; ana] tubercles prominent 15 15a. (14b) Respiratory histoblast with 6 filaments; larvae reddish. S. slossonae 15b. Respiratory histoblast with 10 filaments; larvae yellow. S. dixiense 16a. (13b) Respiratory histoblast filaments 4 in number; gular notch longer than wide. S. species No. 1 16b. Respiratory histoblast filaments number more than 4; the gular notch not longer than wide 17 17a. (16b) Respiratory histoblast with 12 filaments; the anal tubercles are large, prominent; a medium-sized larva, 6 mm long. S. congareenarum 17b. Respiratory histoblast with 16 filaments; the anal tubercles are absent or inconspicuous; a large larva, 3-9 mm long . . S. vittatwn

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-74A k ey to the pupae of the black flics of Florida . (No pupae of Cnephia species No. 1 have been observed.) la. Cocoon a loose mass of silk, indistinct shape; strong dorsal hooks at the posterior end of the pupa. C. ornithophilia lb. Cocoon a distinct slipper or pocket shape, strong dorsal hooks absent . " 2 2a. (lb) Cocoon with large anterior, lateral apertures 3 2b. Cocoon uniform without large lateral apertures 10 3a. (2a) Pupa with 6 respiratory filaments on each side; cocoon antero-ventrally completely joined (Fig. 75) S. notiale 3b. Pupa with more than 6 filaments 4 th 4a. (3b) Pupa with 6 filaments rising off a strong basal 7 one (Fig. 48) S. haysi 4b. Pupa with more than 7 filaments 5 5a. (4b) Pupa with S filaments with four pairs of ?.. ... S. taxodium 5b. Pupa with more than S filaments 6 6a. (5b) Pupa with 9 filaments in the pattern 2, 2, 2 and 3 from the dorsal S. lakei 6b. Pupa with 10 filaments 7 7a. (6b) Pupa with 9 filaments rising from a thick basal 10 one (Fig. 60) S. jonesi 7b. Pupal respiratory organ lacking a strong basal filament .... 8 8a. (7b) Pupa with 10 respiratory filaments which rise as 5 pairs, the petiole for filaments 7 and 8 (from the dorsal) noticiably longer

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-75and stouter than the other petioles (Fig. 12) S. dixiense 8b. Pupa with 10 filaments, some of which rise in pairs others in triplets 9 9a. (8b) The lower more ventral respiratory filaments on long petioles, filaments long S. nyssa 9b. Lower filaments rise close to the base, petioles short, filaments short -5. jenningsi 10a. (2b) Pupa with 4 respiratory filaments S. species No. 1 10b. Pupa with more than 4 filaments 11 11a. (10b) Pupae with 6 filaments in the respiratory organ 12 lib. Pupae with more than 6 filaments 14 12a. (11a) Pupal cocoon with concave anterior edge (in lateral view); respiratory filaments rising from long petioles (Fig. 28)5. slossonae 12b. Pupal cocoon with convex or straight anterior edges; petioles short 13 13a. (12b) Respiratory filaments widespread, base of filaments 5 and 6 sharply separated from base of 3 and 4, filaments long (Fig. 115). S. verccundum 13b. Respiratory filaments noc widespread, bases about equidistant, filaments short (Fig. 10S) S. tuberosum 14a. (lib) Puna with 8 respiratory filaments; cocoon with rough texture. S. decorum 14b. More than 8 filaments . . 15 15a. (14b) Pupa with 12 respiratory filaments; cocoon with long anterior

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-76dorsal projection (Fig. 35) S. concaveenavum 15b. Pupa with more than 12 filaments; cocoon lacking dorsal projection 16 16a. (15b) Pupa with 16 filaments in 8 pairs S. vittatwn 16b. Pupa with more than 20 filaments in pairs and in 3's .S. mevidionale A key to the adult male black flies of Florida. (Adapted and modified from Stone and Snoddy, 1969) la. Second tarsal segment of the hind leg without a pedisulcus but a shallow depression may be present; radius vein with hair' dorsally on the basal segment; large basal cell present . . .C. ovnithophitia lb. Second tarsal segment of the hind leg with a deep pedisulcus; base of radius with or without hair dorsally; iarge basal cell absent. (Simuliiurn) . . .2 2a. (lb) Basal portion of radius with hair dorsally . S. eongareenarum 2b. Basal portion of radius without hair dorsally 3 3a. (2b) Distimere short, stout, with 3 or more teeth ...£'. vittatwn 3b. Distimere long, with 1 tooth or none 4 4a. (3b) Distimere with a rounded lobe on the inner margin near the base 5 4b. Distimere without a lobe or basal projection 6 5a. (4a) Distimere with basal lobe bearing a number of stout, spinelike setae S. tuberosum 5b. Distimere with basal lobe bearing fine hairs only. . . S. slosscmae

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-776a. (4b) Ventral plate in ventral view broadly rounded. S. mevidionale 6b. Ventral plate in ventral view more narrow, compressed from the ,, 7 sides 7a. (6b) Ventral plate with basal arms bearing distinct lateral projections; posterior third of scutum shiny with indistinct hairs. 8 7b. Ventral plate with basal arms without distinct lateral projections; posterior quarter or less of scutum shiny with some strong erect hairs U 8a. (7a) Ventral plate in ventral view with median portion longer t-.han wide and parallel-sided or nearly so; scutum with silver spots narrow, oblique; the dark area between the spots broadens anteriorly. S. dixiense S. jenningsi S. notiale 8b. Ventral plate in ventral view with median portion not longer than wide and widened toward the end; anterior silver spots and intervening dark area variable 9a. (8b) Scutum with large anterior silvery spots each extending about one-third the distance across the front of the scutum; the dark area between the spots narrows strongly to the anterior margin. .S. jonesiS. taxodium 9b. Scutum with smaller silvery spots, the dark area between the spots is broad, converges little, and is narrowest about midway along the length of the spots s haySV S. lakei S. nussa

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-7810a. (7b) Ventral plate in ventral view narrow, V-shaped, middle region tapers almost to a point; with a ventral keel. . S. decorum 10b. Ventral plate broader, not as strongly compressed in the middle section; ventral keel absent S. vereeundum A key to the adult female black flies of Florida . (Adapted and modified from Stone and Snoddy, 1969) la. The second hind tarsal segment lacks a pedisulcus, although there may be a slight depression; the basal cell is present and the basal portion of the radius bears setae dorsally .... C. ornithophilia lb. The second hind tarsal segment with a deep pedisulcus; large basal cell absent and the basal portion of the radius bears or lacks setae dorsally (Simuliuni) 2 2a. (lb) Radius vein with hair dorsally on the basal section. S. eongareenarum 2b. Radius without hair dorsally on the basal section 3 3a. (2b) Tarsal claw with a prominent basal tooth . > 4 3b. Tarsal claw simple, without a prominent basal tooth 5 4a. (3a) Frons gray pollinose S. meridional e 4b. Frons shiny black S. slossonae 5a. (3b) Frons and terminal abdominal tergites shiny black or dark brown 6 5b. Frons and terminal abdominal tergites at least lightly pollinose. 10 6a. (5a) Subcostal vein with a row of hairs ventrally; scutum subshiny 7

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•796b. Subcostal vein without hairs ventrally; scutum shiny 8 7a. (6a) Fore tibia with a narrow gray-white patch not more than one third the width of the tibia; inner margins of ovipositor lobes fairly straight 5". tuberosum 7b. Fore tibia with a brilliant white, patch which covers one half or more of the tibia; inner margins of the ovipositor lobes concave enclosing an oval area S. vevecundum 8a. (6b) In anterior view the scutum displays a pair of fairly distinct rounded pollinose spots with a darker area of the scutum in between; hairs on the stem vein are dark brown to black .... S. jenningsi S. notiale 3. nyssa S. taxodivjn 8b. In anterior view the scutum is not pollinose or the pollinosity • occurs as a diffuse area along the front and sides 9 9a. (8b) Clypeus about as wide as long S. jonesi S. lakei 9b. Clypeus longer than wide S. dixiensc S. hay si 10a. (5b) Scutum silvery gray with dark brown markings; abdomen with a black and light gray pattern S. vittatum 10b. Scutum uniform brownish gray without contrasting dark brown markings; abdomen black with thin gray pollinosity without a pattern. 5. deoovum

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-80Introduction to the Individual Species Sections The following sections deal specifically with the black fly species found in Florida. References are listed below each species name which provide synonyms for the currently accepted name and additional sources of descriptions and figures. The descriptions which follow the brief taxonomy portions are intended to point out outstanding or diagnostic characters and are accompanied with photographic illustrations. Under "Florida Observations" below the heading "Stream Parameters" a summary is given of the dimension, pH, temperature, and velocity figures for streams which contained each particular species. lis the. section on Florida collection records the numbers immediately following the county names refer to individual collection sites idenr.iiiej in the Appendix. The rode for the collection records is as follows: S refers to small, very young larvae with no or weakly developed respiratory histoblasts; M refers to medium-aged larvae with distinct but white histoblasts; •L refers to large, mature larvae with dark histoblasts; ?_ refers to full pupae or pupal exuviae; C refers to cocoons; and A refers to male or female adults. The majority of records refer to specimens the author personally collected; in those instances where specimens were gathered by other individuals, the collector is noted. On the individual species maps when a record is included with the only locality information being the county the mark for that record on the map is placed in the center of the county. Immature, very young larvae lacking well-developed respiratory histoblasts in the subgenus Ftiovl.nxKloros are difficult if not impossible ro differentiate as are the adult females. Records for the Phostevodor-os species are based primarily on mature larvae and pupae. Only at locations where

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-81essentially a single Phostevodoros species was found to exist are records for earlier stages included. Records for undetermined species are listed as S. (Phosterodoros ) spp. in the Manitoba trap results (Table 4). Dr. E.L. Snoddy kindly examined immature and adult specimens of S. jenningsi, S. lakei and S. taxodium from Florida and confirmed my identifications. Cnephia (Cnephia) ovnith.oph.ilia Davies, Peterson, and Wood Cnephia ornithcphilia Davies, Peterson and Wood, 1962, Proc. Entomol. Soc. Ontario 92: 102 (female). Cnephia omithophilia — Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 25 (female, larva). Taxonomy . Davies et al. (1962) first described a female of this species. The holotype was collected from a blue jay at Algonquin Park, Ontario and was deposited in the Canadian National Museum. Paratypes are located in the U.S. National Museum. Stone and Snoddy (1969) indicated the male of C. ornithophilia was not known and that the larva and pupa were apparently indistinguishable from those of C. pecuarum. Description . The larva is large, about 8 mm long with a grayish brown abdomen and a light yellow brown head capsule. The cephalic spots are dark with the median groups of about 20 forming one long continuous row (Fig. 18). The submentum is convex apically and bears small teeth. The gular notch has fairly parallel sides, is broadly rounded anteriorly and extends less than 1/3 the distance to the submental teeth (Fig. 10). The cephalic fans each contain about 60 rays with long hair-like spines. The anal tubercles are absent.

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-82I Figure 18. The head spots of a larva of C. ornithophilia. Figure 19. The pupal exuviura and cocoon of C. ormithovhilia.

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-83The pupa bears strong dorsal posterior hooks and is located in an irregular, loosely woven cocoon. The respiratory organs each contain about 30 filaments (Fig. 19). This is believed to be the first description of the male. The wings are 3.75-4 mm long. There are hairs dorsally on the base of the radius. The first segment of the flagellum is the longest, about twice the length of the other segments. The scutum is dull brownish black with numerous short golden hairs. The scutellum is reddish brown and bears long dark hairs. The legs are almost uniform yellow brown. The second hind tarsal segment lacks a pedisulcus. The abdomen is dull brownish black dorsally and lighter gray or light brown ventrally. The ventral plate in ventral view is broad and the tapering distimere which is almost equal in length to the basimere has a single tooth at its apex as in Fig. 15. Plesiotype: Male, with associated pupal exuvium and cocoon, NW 23rd Ave. and NW 83rd St., Gainesville, Alachua Co., Florida, 2 February 1975 (Pinkovsky) , to be deposited in the U.S. National Museum. The wings of the female are about 4.5 mm long. The base of the radius is covered with setae and a basal cell is present on the wing. The frons bears short yellow hair. The pedisulcus is essentially absent and the prominent tooth on the basal claw has convex margins (Fig. 20). The apex of the abdomen is shiny. The genital fork has widespread narrow arms as in Fig. 21. Distribution . Stone and Snoddy (1969) indicate C. omitkophilia occurs in Louisiana, Mississippi, South Carolina, Virginia, and Ontario. Tarshis and Stuht (1970) report this species from Maryland. Life History . Stone and Snoddy (1969) mention that C. ovv.ithophilia

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-84-• Figure 20. Tarsal claw of a female of C. ornithophilia. I Figure 21. Genital fork and terminalia of a C. ornithophilia female.

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-85is probably univoltine. Eggs collected in stream bottom samples and placed in laboratory rearing tanks yielded first instar larvae three days later (Tarshis, 1973). Larvae have been collected between 11 January and 18 March in South Carolina and adult females were reared on 6 February from pupae; females have been collected from 6 February to 1 May in Louisiana and Mississippi (Stone and Snoddy, 1969). Tarshis (1973) working in Maryland found larvae from 14 November to 29 April, pupae from 11 January to 22 April and adults from 2 to 17 March. Tarshis and Stuht (1970) captured adult C. ormithophilia between 28 February and 1 April in emergence cages over a stream in Maryland. When first to third instar larvae were collected and reared in the laboratory in aerated water at 15-22°C pupae appeared after 2 to 21 days and adults emerged 4 to 23 days after introduction of the larvae (Tarshis, 1973). Tarshis (1973) found adults emerged as soon as 2.5 hours after pupation [very difficult to believe] and that the number of males emerging exceeded the number of females emerging only on the first day. Ecolog y. Tarshis and Stuht (1970) found massive numbers of larvae on leaves, rocks, and twigs in a stream that flowed .153 to 1.0 m/sec (.49-3.3 ft/sec) from a pond through an oak and willow woods. The stream was .92-1.53 m (3-5 ft) wide, 2.5-20 cm (1-8 in) deep and had a pH reaction between 6.9 and 7.4. In a similar stream that was slightly deeper and 1.22-3.66 m (4-12 ft) wide, larvae were collected at water temperatures of .5-15°C (Tarshis, 1973). Tarshis (1973) mentioned that pupae easily suffered injury and that development to adults did not occur when pupae were removed from their substrate. Larvae of C. mutata. P. gibsoniy S. decorum, S. tuberosum, S. venustum, and S. vtttatum were collected with C. ornithophilia in a pond runoff stream where

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-86C. ornithophilia predominated (Tarshis and Stuht, 1970). Habits . Tarshis and Stuht (1970) suggest that C. ornithophilia oviposits on the stream surface and the eggs sink to the stream bed. Bennett (1960) in Ontario recovered C. ornithophilia (as Cnephia "U") from jays, hawks, robins, sparrows and other woodland birds. Tarshis (1972) fed C. ornithophiliaon geese and ducklings in the laboratory and transmitted L. simondi to ducklings by the bite of the fly. Tarshis (1973) did not collect any C. ornithophilia from ducklings exposed on the banks of a stream containing larvae and pupae of the black fly and suggests that the target hosts were inappropriate or perhaps that the flies were absent at ground level and concentrated in the canopy. Florida o bservations . Stream Parameters Width Depth pH Temperature Velocity Mean: 2.57m 23.64 cm 4.7 14.7°C (58.6°F) .45m/sec (1.47 ft/sec) Min: .3 1.3 3.6 8.3 (47) .23 (.75) Max: 11 90 6.6 24.4 (76) 1 (3) Cnephia ornithophilia is reported here for the first time from Florida. Both Stone (1965) and Stone and Snoddy (1969) mention that. C. pecwxrim occurs in Florida. Key characters, such as the shape of the tooth on the tarsal claw of the female, on specinens I obtained in Florida differed from those of C. pocuavum. In the U.S. National Museum I located one adult Cnephia specimen from Florida, listed for Site 204 below, which was a male that had been labeled as C. pcauarum but C. ornithophilia was penciled in underneath. Adult and immature specimens which I submitted flora five collections representing two west Florida (Holmes and Liberty) and one east Florida. (Alachua) counties were

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-87identified by G.E. Shewell of the Canadian Biosystematics Research Institute as C. ornithophilia. Cnevhia ornithophilia was collected from 31 sites in 14 counties in Florida (Fig. 22). Most of the records represent collections of only a few, usually young larvae with small, pale histoblasts, but the larger overall body size compared to other species and distinctive head spots and gular notch are discernible even in young larvae. Larvae, pupae and associated adults were obtained at three sites — 28, 119 (Fig. 23), and 147). This species was first collected in the streams on 28 October and last collected on 18 April. Most collections occurred and the largest populations were encountered during December through March. Collections of C. ornithophilia at a number of sites were made during one winter month and the species was not found again until about the same time the following year. It appears that C. ornithophilia is univoltine in Florida. This species occurs in cool, fairly shallow flows, usually at most a few meters wide, most frequently with a velocity of .305-. 61 m/sec (1-2 ft/sec) and a pH which normally is below 5. Immatures were also collected at Lebanon Station (Site 139) and Yellow Water Creek (Site 70) where the pH approaches neutrality. At most sites there was considerable vegetation especially trailing grass in the stream to which the immatures were attached. At Blues Creek (Site 2) and Site 28 sparce vegetation was present. Site 28 originated in a small swamp just upstream, was about .5 m wide, 10 cm or less deep and tumbled down a root-filled and rocky bed through woods. In January a heavy population of C. ornithophilia covered rocks, dead tree leaves, pine needles, and a few blades of trailing grass. Masses of 40 or more larvae encircled twigs about .6 cm in diameter in the current. Most of the

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j -a* Figure 22. Collection locations for C. ormithophilia in Florida.

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-89Figure 23. Site 119, Gum Creek, a stream inhabited by C. ornithophilia.

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-90streams in which C. ornithophilia was located ceased flowing sometime during the year. At Site 99 at the Alapaha River C. ornithophilia was only collected when the river rose out of its normal channel and poured alongside the highway in flows into wooded areas. In the streams with C. ornithophilia were found nine other black fly species (Table 3 ) . Most frequently S. slossonae and S. tuberosum, both with 32 associations, were collected with C. ornithophilia. The next most common associates were S. congareenarum and S. vereoundum which have seasonal occurrences that are similar to that of C. ornithophilia. Swollen larvae infected with white masses of thousands of small white spheres were observed on at least five occasions. Usually these larvae were 4-6% of the total number present. While dissecting adult females I observed that the salivary glands were large compared to those of S. slossonae and S. congareenarum and similar to the findings of Bennett (1963b), I found the glands lacked the golden pigment typical of most ornithophilic species. The eggs of C. ornithophilia were noted to be unusual also. Instead of being subtriangular as with the Simulium species the eggs of C. ornithophilia (.3 mm long and .1 mm wide) tapered to a point at both ends like a spindle. Immatures collected with their vegetation substrates from Site 119 on 31 December and reared in aerated water continued to yield emerging adults until 12 January. From larvae and pupae collected at Site 28 and reared in the laboratory 26 male and 25 female C. ornithophilia were obtained. Adult females were captured in a Malaise trap at Site 2 at the IFAS Horticulture Unit during March. The record of Site 12 below is that of a C. ornithophilia female captured with a S. slossonae female feeding on

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-91chickens. Six C. omithophilia fed on turkeys in the lab but did not vector L. smithi. One female that emerged on 2 January was maintained on moist cotton and raisins in a holding carton and died 16 days later. Another female emerged on 4 January, fed on an infected turkey on 11 January, fed on clean birds on 14 and 19 January, and died on 25 January, 21 days after emergence. A third female fed twice on turkeys and died on 26 January or 22 days after she emerged. Florida collection records for C. omithophilia . Alachua Co. 1) 1976: 21 Jan (S). 2) 1974: 2 Jan (S,M,L), 14 Dec (S) ; 1975: 10 Jan (S) , 19-20 and 21-23 March (A G.B. Fairchild) . 6) 1974: 2 Jan (S,M), 7 March (S) . 8) 1975: 10 Jan (S,M). 12) 1976: 22 Feb (A H. Davis). 14) 1974: 7 March (S). 21) 1974: 2 March (S,M); 1976: 21 Jan (S,M). 23) 1975: 24 Jan (S). 28) 1975: 28 Oct (S), 9 Nov (S) ; 1976: 30 Jan (S,M,L,P,A). Bradford Co. 43) 1975: 12 Jan (L) ; 1976: 31 Jan (S,M). 44) 1974: 4 Jan (S); 1976: 31 Jan (S) . 48) 1975: 26 Jan (S,M). Dixie Co. 68) 1975: 17 Jan (S). Duval Co. 2°) 1974: 4 Jan (M,L,P); 1976: 14 Feb (S,P). 12) 1974: 5 Jan (M); 1976: 14 Feb (S) . Hamilton Co. 99) 1975: 1 Feb (S,M); 1976: 14 Feb (S) . Holmes Co. 119 ) 1975: 19 Jan (S,M,L,P), 28 March (S,M,P,A), 30 Dec (S,M,L,P,A); 1976: 18 April (M) . Leon Co. 131 ) 1973: 17 Dec (S,M,L); 1975: 17 Jan (S) , 29 Dec (S) . 134 ) 1973: 19 Dec (S). Levy Co. 135) 1973: 29 Dec (S). J.39) 1975: 21 Dec (M,L). Liberty Co. 142) 1975: 18 Jan (M) . 147) 1973: 18 Dec (S) ; 1975:

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-9218 Jan (S,M,L,P,A), 30 Dec (P). Nassau Co. 159 ) 1974: 4 Jan (S,M,L); 1975: 12 March (M) . 160 ) 1975: 12 March (M) . 161 ) 1976: 14 Feb (L) . 162 ) 1975: 12 March (P) ; 1976: 14 Feb (S,M). Okaloosa Co. 168) 1973: 13 March (S,L — K. Tennessen) . Seminole Co. 204 ) 1960: 16 Nov (AE.G. Jay). Taylor Co. 211 ) 1975: 17 Jan (M) . Union Co. 217 ) 1974: 5 Jan (M) . Simulium (Byssodon) mevidionale Riley Simulium mevidionale Riley, 1887, Rep. of Entomol. , U.S. Dep. Agr. Rep. for 1886: 513 (female). Simulium oacidentale Townsend, 1891, Psyche 6: 107 (female). Simulium tamulipense Townsend, 1897, J. N.Y. Entomol. Soc. 5: 171 (female) . Simulium f orb e si Malloch, 1914, U.S. Dep. Agr., Bur. Entomol-. Tech. Ser. 26: 63 (female, male, pupa). Simulium mevidionale — Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 28 (female, male, pupa, larva). Taxonomy . Of the larva, pupa, male, and female in the original description by Riley (1887) only the female was not misidentif ied (Stone and Snoddy, 1969). Dyar and Shannon (1927) indicate the type locality for S. mevidionale is probably Lake View, Mississippi and that type material is located in the U.S. National Museum. Stone and Snoddy (1969) mention that S. mevidionale as recognized in the United States may actually refer to a group of sibling species.

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-93Descrlption . The larva is described by Stone and Snoddy (1969) as possessing large ventral tubercles, antennae which lack hyaline bands, and respiratory histoblasts which in the normal coiled condition are only slightly concave along the posterior border. The cephalic apotome bears a central posterior light spot surrounded by a fulvous ring with two dark pigmented rays projecting and diverging anteriorly. The pupa is about 3 mm long and is situated in a well-constructed, slipper-shaped cocoon that projects forward noticeably along the ventral half. The respiratory organs each consist of about 25 filaments (Fig. 24). The male as described by Malloch (1914) (as S. forbcsi) and Stone and Snoddy (1969) is 2-2.5 mm long and possesses a velvety black to dark reddish brown scutum which lacks silvery spots. The abdomen is velvety black with a yellowish venter. The ventral plate is broad in ventral view, lacks marginal denticles and is not deeply notched. In end view the ventral plate displays only a very small median lobe. The female wing is 2-2.5 mm long. The female is gray in appearance with a gray, poliinose frons and a gray scutum with three dark longitudinal lines (Fig. 25). The fore coxae are dark. The genital fork has a fairly thick stem and wide arms with dark, prominent, apical, dorsal projections (Fig. 17). Distribution. Dyar and Shannon (1927) listed S. meridionals (as S. pad dentate) from Jacksonville, Florida. Stone (1952) reported S. mcridionale from a few sites in Alaska. Shewell (1957) recorded S. meridional?, as an Austral and southern Boreal species from the northern transition regions of Canada. Snow et al. (1958) captured one female. S. mcridionale at Sugar Tree, Tennessee. Stone and Snoddy (1969)

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-94Figure 24. The pupa and cocoon of S. mevidionate . Figure 25. The scutum of a S. meridionale female.

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-95indicate the distribution is from Alaska to Indiana and south to California, Florida, and Mexico. Life History . Stone and Snoddy (1969) indicate that the eggs of S. meridionals overwinter. Anderson and Dicke (1960) found adults could be collected from late May to late October and suggest there are at least four generations each year. Larvae matured in three weeks and pupae matured in three to four days in water temperatures 20°-24.4°C (6£-76°F) (Anderson and Dicke, 1960). Stone and Snoddy (1969) report at least four generations a year in Alabama with adults being collected from 16 March to 24 December. Ecology . Shewell (1958) stated that S. mevidionale is clearly a big river species. Tucker (1920) reported that adults appeared during times of high water and overflows of the Mississippi River. Townsend (1891) reported that S. mevidionale (as S. oaaidentale) adults appeared as the Rio Grande River rose during May and June. Anderson and Dicke (1960) found immature stages in the Mississippi and Wisconsin rivers commonly attached to the sides of small rocks or beneath grass blades 5-15 cm (2-6 in) deep in currents usually less than .3 m/sec (1 ft/sec). Habits . Anderson and Dicke (1960) collected adults up to 24 km (15 mi) from known breeding areas. The common name of S. mevidionale is the turkey gnat (Blickenstaf f , 1970). Anderson and DeFoliart (1961) found S. mevidionale fed on white and bronze turkeys, chickens, pheasants, doves, and starlings with more flies being attracted to caged avian hosts placed at the tree canopy level than at ground level. Skidmore (1931) showed that S. mevidionale was a vector of L. smithi to turkeys. Edgar (1953) reported a serious decline in egg production in chickens troubled by a spring outbreak of S. mevidionale in Alabama.

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-96Si.nruli.im meridionale was reported heavy and damaging to chickens in five Alabama counties in 1976 (U.S.D.A., 1976). Anderson et al. (1961) recovered eastern encephalitis virus from a pool of unengorged 3. meridionale in a turkey brooder house and suggested that this species might serve as a biological vector. Townsend (1891), Anderson and DeFoliart (1961) and DeFoliart and Rao (1965) report that S. meridionale feeds on man. DeFoliart and Rao (1965) observed a greater tendency in S. meridionale to bite man after the flies had been exposed to a period of cool temperatures and suggested that S. meridionale might transmit encephalitis to humans. Florida observations . Based primarily on adult collections, S. meridionale is recorded from 10 locations in 5 Florida counties (Fig. 26). The single record from Duval County refers to specimens collected years ago in Jacksonville by Mrs. A.T. Slosson and observed at the U.S. National Museum. Adults have been captured on the wing from 15 April to 26 May in counties bordering the Apalachicola River which is the only currently known breeding location of the species in Florida. Pupae of S. meridionale and two young, not positively identifiable larvae were collected on 17 April 1976 along the Apalachicola River at Richbourg's Landing (Site 145). The river was just beginning to recede from its spring flooding and had dropped about 1.83 m (6 ft) exposing rocks, some roots and sparse vegetation. At the collection site steep 9 m (30 ft) high banks of crumbling soil and rock were separated from the river by a strip of concrete and pebbles a few meters wide. A thick root sticking up vertically a short distance out of the water was retrieved and immatures were found on it from the apex to 55 cm (almost 2 ft) down. One large clump (1.8 x .8 m) of green grass-like vegetation with blades 15 cm (6 in) long and .9 cm (about 3/8 in) wide that was exposed on the

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-97rj JJ&' Figure 26. Collection locations for S. meridionals in Florida.

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-98bank about 1 m out of the water harbored a large population of, mainly, 5. meridionale pupae as did one small exposed bush. Other trailing tree roots yielded immatures but none were found on the pebbles or concrete in or out of the flow. The river was silty yellow brown, extremely wide and apparently very deep beyond the concrete strip. The current was about .61 m/sec (2 ft/sec), the water temperature was 21.5°C (70.5°F) and the pH was 6.5. Collected with the S. meridtonale immatures were pupae of S. jonesi. Some adult S. meridtonale were observed flying about the collection area. Only pupae and adult females of S. meridtonale have been collected in this research. The record for Site 53 is for S. meridional? adults at Blountstown which were reported to be causing deaths to young chicks. A number of reports of pouiury and human suffering were received during collecting trips in Liberty and Calhoun counties during the April 1975 and 1976 S. meridionale mass emergences. Severe swelling and pain reactions to the bites of the. bull flies, as the S. meridionale adults were called, were reported. The author found having twenty to thirty flies circling around the head, entering one's ears and crawling on one's neck and clothes slightly annoying, but he was never bitten. The collection records indicate a single generation for this species in Florida. On dissecting wild -caught S. meridionale females, in the ovarioles next to well-developed eggs were noted small bodies which may have been small, developing eggs, possibly an indication of two ovarian cycles. One S. meridionale which fed on a turkey infected with L. smithi was observed to feed on a clean turkey three days later. The clean bird became positive for L. smithi thus incriminating S. meridionale as a

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-99vector of the disease in Florida. Wild caught flies were kept alive in the lab on water and raisins for a maximum of ten days. Florida collection records for S. meridionale . Calhoun Co. 51) 1975: 19 April (A). 53) 1973: 10 May (A). 52) 1975: 19 April (A). Duval Co. 71) (no date): (A -Mrs. A.T. Slosson) . 1 Franklin Co. 80) 1976 Jackson Co. 122 ) 1973 Liberty Co. 145 ) 1976 2-3 April (A-.G.B. Fairchild). 26 May (A W.W. Wirth) . 17 April (P,C). 149 ) 1975: 19 April (A). 150 ) 1975: 19 April (A); 1976: 17 April (A). 154 ) 1957: 15 April (A F.S. Blanton) ; 1966: 20 May (A H.V. Weems). Simulium (Byssodon) slossonae Dyar and Shannon Simulium slossonae Dyar and Shannon, 1927, U.S. Nat. Mus. Proc. 69(10): 34 (female, male). Simulium slossonae — Underhill, 1944, Va. Agr. Exp. Sta. Bull. 94: 21 (female, male, pupa, larva). Simuliwn slossonae — Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 29 (female, male, pupa, larva). Taxonomy . Dyar and Shannon (1927) briefly described the female of S. slossonae and also the male genitalia. The type locality is given as Biscayne Bay, Florida. The type male and seven paratype females were deposited in the U.S. National Museum (Dyar and Shannon, 1927). Underhill (1944) provided more detailed descriptions of the larva, pupa, male and female. Description. The larvae are 4.5 to 5 mm long with reddish-brown

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-100mottling and banding on the abdomen and thoracic region. The head spots are dark and distinct on a more pale head capsule. The posteriorlateral pair of spots on both sides are distinctly curved (Fig. 9 ). The gular notch is elongate, widest at three-fourths the way along its length anteriorly and tapers to a broad point (Fig. 27). There are 48 to 56 rays in each cephalic fan. The anterior arms of the anal crosspiece appear long. The anal lobes are large and cone-shaped. The pupa is about 2.5 mm long in a slipper-shaped cocoon with distinctly concave anterior margins and a short, broad, anterior, middorsal projection. The six, slender respiratory filaments arise in three pairs, lie close to each other, and have long petioles one-fifth to one-fourth the length of the filaments (Fig. 28). The male is about 2.5 mm long and possesses a dark gray shiny scutum with black velvety areas centrally across the scutum which form an anterior triangle. The ventral plate of the male in end view is broadly rounded; the distimere is curved inward, rather flat and bears a lobe with hairs, basally (Fig. 29). The female is about 2.5 mm long and is shining black with a shiny black frons. The abdomen bears three wide velvety black bands dorsally on the first few abdominal segments and more distally displays glistening black tergites. The female lacks setae on the underside of the subcosta and the base of the radius is bare dorsally. Each tarsal claw has a prominent basal tooth. The genital fork has thick arms with ventral projections which curve in toward each other forming a space like a U with a constricted opening (Fig. 30). Distribution . Dyar and Shannon (1927) list S. slossorxte from Biscayne Bay, Florida, and South Carolina. Underhill (1944) found this

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-101h Figure 27. Gular notch of a 5. slossonae larva. Figure 28. The pupa and cocoon of S. slossonae.

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-102m & i Figure 29. Terminalia of a male of S. slossonae. Figure 30. Terminalia of a female of S. slossonae.

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-103species in the tidewater section of Virginia. Snow et al. (1958) do not list S. s lossonae among the species occurring in the Tennessee River Valley. Stone and Snoddy (1969) list the additional states of Alabama, North Carolina, Georgia, Mississippi, and Texas for the distribution of S. slossonae. Life Cy cle. Larvae, pupae, and adults were collected as early as February in Jasper County, South Carolina (Jones and Richey, 1956). In Virginia S. slossonae was first found in collections made during March and last found during November (Underhill, 1944). Garris et al. (1975) found immatures of S. slossonae throughout a May through October collecting period. Jones and Richey (1956) concluded that the life cycle of S. slossonae was shorter than one month based on the discovery of many larvae and pupae, eight and twenty-one days, respectively, after a rain stimulated flow in streams previously devoid of black flies. Garris ct al. (1975) state that S. slossonae is a multivoltine species present al] year. In Alabama it was found from January through December except during June and September (Stone and Snoddy, 1969). Ecology. Underhill (1944) mentioned that streams in Virginia inhabited by S. slossonae were .92-3 m (3 to 10 ft) wide, 20.3-38 cm (8 to 15 in) deep, usually had sandy-mud bottoms and were sJow flowing with the swiftest portions reaching about .46-. 61 m/sec (1.5-2 ft/sec). Jones and Richey (1956) found larvae in partially shaded sections of streams on tape grass and record maximum populations of 26 larvae per 2 2 6.5 cm (1 in ) of leaf surface. Significant plant growth and shade were typical of streams containing S. slossonae in Virginia (Underhill, 1944). Stone and Snoddy (1969) state that S. slossonae occurs in the swamp rivers of the South.

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-104Hab its . Underhill (1944) reports that S. slossvnae feeds on turkeys. Wehr (1953) first showed S. slossonae could transmit L. smithi. experimentally, by intramuscular injection. Jones and Richey (1956) succeeded in transmitting L. smithi to a previously uninfected turkey by the bite of a S. slossonae female that had fed five days earlier on an infected bird. In South Carolina S. slossonae was captured from turkeys and humans but did not bite humans (Jones and Richey, 1956). Noblet et al. (1975) found that peaks in the adult populations of S. slossonae occurred during lets July, late September, and late October, that the numbers of S. slossonae feeding on turkeys declined in November, and that few adult S. slossonae were present from mid-November until midFebruary. Moore and Noblet (1974) report that S. slossonae commonly flies four miles after engorgement usually following stream courses toward swamps and mention adults may travel up to eight miles. Florida observa t ions . Stream Parameters Width Depth pH Temperature Velocity Mean: 3.47 m 31.68cm 4.5 21.3°C (70.3°F) .44 m/sec (1.45 ft/sec) Min: .076 1 3.5 8.9 (48) .12 ( .4) Max: 31 200 6.9 28.9 (84) 1.02 (3.33) Stone and Snoddy (1969) state that S. slossonae is abundant in Florida and Davis ct al. (1957) mention it was the most prevalent species encountered in Florida. I found that S. slossonae occurs in some streams all year long in Florida. The collection records listed below indicate that S, slossonae is the most widely distributed species in the State and has been collected at one time or another from 113 sites in a total of 46 counties (Fig. 31). Tt occurs in many parts

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-105rJ J&** Figure 31. Collection locations for 5. slossonae in Florida.

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-106of west Florida, along the Gulf coast (Dixie, Levy Co.'s) where other species such as S. tuberosum are missing, is well established in north central Florida and is found as far south as just west of Lake Okeechobee. The most southern spot at which S. stossonae was found during this research was in Fisheating Creek which flows through cypress swamps in Glades County. The record from Dade County refers to material collected by Mrs. A.T. Slosson, probably around 1900 at a time when the Miami River had rapids. The distribution of S. stossonae most likely extends into other counties and a greater range will probably be demonstrated by later investigators. As indicated by the stream parameters above S. stossonae was found in streams which on the average were fairly small and slow moving, with a mean velocity below .46 m/sec (1.5 ft/sec). These flows often exhibited a red brown or tea color and a low pH reaction in the vicinity of 4.4-4.5. In addition S. stossonae preferred streams with considerable stream vegetation and without excessive shading. Sites with most or all of these characteristics which were inhabited by S. stossonae most of the year and had large populations of immatures during certain periods included Sandy Hatchet Creek (Site 1) , a tributary to Hatchet Creek (6), Site 8, Lcchloosa Creek (21), Sites 43 (Fig. 32), 44, and 119, the stream from Lake Melrose (185) and Turkey Creek (216). Site 211 one mile west of the Fenholloway River is a location where S. stossonae was found year round in water of very low pH (3.5-4.0). Other streams (Sites 23 and 48) are very temporary in nature and clogged with grass and other green vegetation but when flowing in August and September contain good populations of S. stossonae. Immatures were found attached to all forms of aquatic vegetation including field and

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-107&r '••., %4> "\v . • -'..r^ PH -tlPl WMffl Figure 32. Site 43, Double Run Creek, where S. slossonae immatures were collected.

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-108eel grass, sedges, smartweed, pickerel weed, cattails, and Hydrilla as well as dead leaves, pine needles, twigs and, in lesser numbers, rocks and concrete chunks. The size of S. slossonae populations seemed to be limited more by competition with other black fly species and actual flow stoppage than by increasing summer stream temperatures. Frequently, in streams that continued to run the largest numbers of immatures were located from April to November. In streams such as 43, 119, and 216, with an April to September period of maximum S. slossonae numbers, the initial, increase in S. slossonae populations appeared to be a response to the increasing spring and early summer water temperatures and a lessening of substrate and perhaps food competition by winter and early spring species like S. congaveeyiavw:' and C. ormitkoph-ilia as their populations began to decline. While most streams from impounded water were inhabited by species such as S. decorum or S. vittatwn, two sites were encountered where S. slossonae was the predominant species. At Site 177, a very intermittent flow below a lake in Pasco County, a tremendous population of S. slossonae including many pupae and cocoons was found durivig December on grass, twigs, dead leaves, and concrete. Site 85, a stream from Lake Melrose, flowed continuously and the smartweed, grasses and other vegetation and concrete chunks harbored good populations all year long. S. slossonae occasionally appeared in streams with a neutral or slightly basic pll reaction (Sites 131, 213, 214) however population sizes did not approach those found in more typical S. slossonae streams. S. slossonae was discovered rarely or only in small numbers in streams in Clay County (Sites 56 and 57) where S. iid>erosum and S. jonesi predominated and a similar situation existed in Hatchet Creek (17) and

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-109Sites 18 and 40 where pH and vegetation in most cases, were appropriate for S. slossonae. At Yellow Water Creek (70) S. slossonae was the predominant species in July and September of 1974 and from March through August 1975 maintained strong populations in coexistence with those of S. tuberosum and S. jonesi. In the streams of Florida S. slossonae was associated at least once with eleven other black fly species (Table 3 ) . Simulium tuberosum was most commonly found where S. slossonae occurred. Next frequently found was S. Qonesv followed by S. oongareenarum, S. lakei, and 5. verecundum. S. slossonae was collected with S. vittatum on only fifteen occasions and never with S. decorum, which reflects the more limited distribution of the latter species and generally different habitat preferences in S. slossonae. No mermithids have been observed in the larvae of S. slossonae in Florida. Larvae have been observed swollen with white masses or infected with groups of tiny white spheres, probably microsporidian infections, on a number of occasions at just over a dozen sites. Infections were rarely found in more than 6 or 7 larvae per collection. In samples of 20 or more larvae of the stage infected infection rates ranged from 1.1% (1 of 90) to 7.7% (7 of 91). At Sites 21 and 186 1 of 14 and 1 of 4 medium-aged larvae were infected, respectively. At Sites 119 and 216 large, mature larvae were found with infected abdomens. Adult females were captured in six counties during January through October in Manitoba traps (Table 4 ) . Captures were made from dawn to dusk in winds from 0-16 km/hr, temperatures from 19.4-34°C and relative humidities from 49-100%. Females were also captured in Malaise traps (Site 25). Adults were observed to feed on domestic turkeys in the

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-110field and in the laboratory during most of the year. Si.muli.ton slossonae has been shown to be the primary vector of L. smithi to turkeys in Florida (see Leucocytozoon transmission results section) . The record for Site 12 below is that of S. slossonae captured while feeding on chickens. Adults have been netted from about the head and off the clothes of the author on a number of occasions even when temperatures in the field reached 32°C (90°F) . Adults were netted during October at Sites 2 and 3 where S. slossonae immatures were never or infrequently found. One S. slossonae was captured biting a human on 5 July, 1972, in Gainesville, Florida (5 ). Katherine Sommerman collected an adult in Orlando in 1953 and labeled its actions as "probing". Florida collection records for S. slossonae . Alachua Co. 1) 1973: 2 Nov (S,M,L,P), 14 Nov (S,M,L,P), 7 Dec (S,M,L,P), 13 Dec (S,M,L,P); 1974: 2 Feb (S,M,P), 7 March (S.M.P), 12 April (S,M,L,P), 5 May (M,P), 7 June (S,M,L,P,C), 6 July (S,M,L,P,C), 19 July (A), 20 July (S,M,L,P,C), 29 Aug (S,M,P,C), 28 Sept (S,M,P), 30 Oct (S,M,L,P), 23 Nov (S,M,L,P); 1975: 10 Jan (S,M,L), 12 Feb (M,P,A), 4 April (S,M,L,P), 10 May (P,C), 3 June (P,A), 21 June (S,M,L,P,C), 15 July (S,M,P,C), 25 Sept (S,M,C), 9 Nov (S,M,L,P, C,A); 1976: 21 Jan (S,M,L,P,A). 2) 1974: 2 Jan (S) , 17 Aug (M,P); 1975: 26 May (C), 3 Oct (A). 3) 1975: 3 Oct (A). 5) 1972: 5 July (AD. Young). 6) 1973: 22 Sept (S,P,A); 1974: 2 Jan (S,M), 2 Feb (S,L,P), 7 Mar (L) , 2 April (S,M), 18 May (S,M,L,P,C), 30 June (S.M.L.P), 30 July (S,M,L,P), 14 Sept (S,L,P,C); 1975: 18 May (S,M), 16 Aug (S,M,L,P,C), 8 Oct (S,M,L,P,C), 6 Dec (S,M, L,P,C). 8) 1973: 5 Sept (S), 2 Nov (L,P) ; 1974: 14 Feb (M) , 2 April (S), 16 May (S,M,L,P), 30 June (S,M,L,P), 6 July (A),

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-1113 Aug (S,M,L,P), 22 Sept (S,M,L,P,A), 25 Oct (S,M,L,P); 1975: 10 Jan (S,M), 12 Feb (S,M,L,P,C,A) , 4 April (S.M.L), 18 May (P,C), 1 June (S,M,L,P,C,A), 18 June (S,M,C), 15 July (S,P), 25 July (P) , 31 July (S,M,L,P,C), 30 Aug (S,M,P,C), 30 Sept (S,M,P,C,A), 9 Nov (S,M,A), 14 Dec (S.M.P); 1976: 6 March (S,M). 9) 1973: 5 Sept (S). 10) 1973: 5 Sept (S). 12) 1976: 22 Feb (A H. Davis). 14) 1973: 5 Sept (S,M,L,P); 1974: 7 March (S) , 30 May (S,M,L,P, C,A), 30 July (S,M,L,P), 14 Sept (S.M.L.P). 17) 1974: 1 July (P) , 24 Aug (A); 1975: 1 March (S), 30 April (M) . 18) 1974: 25 May (M), 24 Aug (M); 1975: 14 June (S), 17 July (A). 19) 1975: 17 July (S,M), 22 Oct (S). 20) 1974: 12 Jan (S), 6 July (S,M,P), 29 Aug (S,M); 1975: 18 April (M) , 3 Oct (M,P,A). 21) 1974: 2 March (S,M,L,P), 11 April (S,M,L,P), 18 June (S) , 28 June (S,M,L), 16 July (A), 30 July (S,M,L,P,C), 14 Sept (S,P); 1975: 24 Jan (S,L,P,A), 4 April (S,M), 27 Sept (S,M,L,P,C); 1976: 21 Jan (S,M, L,P,C), 29 Jan (M,P,A). 22) 1974: 28 June (A), 15 Aug (A), 28 Aug (S,M,L), 15 Sept (A), 7 Dec (S) ; 1975: 24 Jan (A), 29 April (S), 7 July (A), 5 Aug (S) , 17 Oct (M,L,P); 1976: 6 March (S,A). 23) 1975: 24 Jan (S,M). 24) 1974: 25 May (M) , 30 May (A), 28 June (A), 16 July (A), 24 July (A); 1975: 18 June (A). 25) 1975: 13 and 15 March (AJ. Glick) . 28) 1975: 9 Nov (L) ; 1976: 26 March (A). 34) 1974: 19 July (A), 17 Aug (A). 36) 1974: 12 April (S,M), 5 May (C) , 6 July (S,M,L,P), 24 Aug (S,M,P,C,A); 1975: 15 Jan (S,M,L,P,C), 4 April (S,M,L,P,A), 22 May (S,M), 22 Oct (S,M,L, P,C,A). Bay Co. 40) 1975: 6 Sept (S) . Baker Co. 41) 1974: 4 Jan (S) , 1 June (S,M,L,P,C), 17 Sept (S,M,L,P,C),

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-1126 Nov (P); 1975: 1 Feb (S), 26 April (S,M,L,P,A), 21 June (S), 17 Aug (S,P), 14 Nov (S,L). Bradford Co. 42) 1973: 6 Oct (S,M,L,P,C); 1974: 4 Jan (S) , 20 July (S.M.P.C), 31 Aug (S,M,L,C), 9 Oct (S,M,L,P,C), 16 Nov (P,C); 1975: 26 Jan (S,C), 15 Sept (S,M,L,P,C), 19 Nov (S,M,L,P,C,A) . 43) 1973: 17 Nov (S,P); 1974: 6 June (P,C), 20 July (S,M,L,P,C), 31 Aug (S,M,L,P,C), 19 Oct (S,M,P,C), 16 Nov (S.M.L.P); 1975: 12 Jan (S,M,L,P), 26 Jan (A), 5 April (S,M,P,C), 6 May (A), 1 July (S,M,L, P,A), 17 Aug (S,M,L,P,C,A), 26 Sept (S,M,L,P,C,A) , 14 Nov (S,M,L,P,C); 1976: 7 Jan (S,M,L,P,C,A) , 31 Jan (S,M,P,C,A), 24 March (A). 44) 1974: 4 Jan (S), 23 Feb (C) , 14 April (S,M,L,P), 10 July (S) , 20 July (S,M,L,P), 31 Aug (S,M,L,P,C,A) , 9 Oct (S,M,L,P,C); 1975: 12 Jan (S,M,L,P,A), 5 April (S,M,L,P,C), 22 May (S,C), 17 July (S,M,L, P,C,A), 26 Sept (S,M,L,P,C,A), 19 Nov (S,M,L,P,C,A) ; 1976: 31 Jan (S). 46) 1975: 26 Sept (S,M,L,P,C). 47) 1974: 20 July (S,P,C), 21 Sept (S,M,L,P,C); 1975: 26 Jan (S), 26 April (S,M,L,P). 48) 1974: 3 Aug (S,M,L,P,A), 21 Sept (M.L.P.A); 1975: 5 April (P) . Calhoun Co. 50) 1975: 19 April (A). Clay Co. 56) 1975: 2 Aug (S,P). Columbia Co. 58) 1974: 21 Sept (S,P). Dade Co. J33) (no date) (A — Mrs. Slosson) . Dixie Co. _67) 1974: 5 Aug (S) . J38) 1975: 17 Jan (P,A), 26 March (S,M,P,C), 23 Aug (S,L,P,C). 69) 1975: 23 Aug (S). Duval Co. 70) 1974: 4 Jan (S,M), 20 April (S,M,L,P,C,A) , 10 July (S,M,P,A), 21 Sept (S,M,L,P,C); 1975: 12 March (S,M,L,P), 4 May (S,M,P,C), 1 July (S,M), 17 Aug (S,M,L,P,C,A) ; 1976: 14 Feb (S,M, L,P). 72) 1974: 5 Jan (M) , 20 April (M,L,P,A), 10 July (S) ; 1975:

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-11312 March (S,M,L,P,A), 4 May (S,M,L,P,C), 1 July (S,M,L,P,C,A) ; 1976: 14 Feb (P) . Escambia Co. 74) 1974: 16 June (S,M), 14 Oct (S) . Flagler Co. 77) 1974: 26 Jan (S) , 25 May (S), 21 Aug (S,M,L,P,C,A) , 7 Dec (P); 1975: 5 April (S,M), 9 July (S,M,P), 31 Oct (S,M,P,C). Franklin Co. 80) 1976: 2-3 April (AG.B. Fairchild) . Gadsden Co. 81) 1974: 15 June (L) . Glades Co. 95) 1974: 11-12 July (A), 26-27 July (S,M,L,C,A); 1975: 17 June (S,M,P,C), 27-28 June (A), 28 July (P,C), 29-30 July (A), 9-10 Aug (S,L,P,C,A), 17-18 Sept (A), 15-16 Oct (A). Gulf Co. 97) 1970: 1-3 May (A W.W.Wirth) . 98) 1970: 3 May (A W.W. Wirth) . Hamilton Co. 99) 1975: 1 Feb (A), 26 April (S,L). 101 ) 1975: 26 April (S,M,P,C,A), 3 Aug (S,P,C,A). Highlands Co. 108 ) 1973: (S,P). 110 ) 1948: 13 July (A R.H. Beamer) ; 1949: Jan (A J.G. Needham) ; 1958: 26 Dec (AS.W. Frost); 1959: 10 Jan (AS.W.Frost), 30 March (AS.W. Frost), 7 and 18 Nov (AS.W. Frost); 1960: 23-24 Feb (AS.W. Frost); 1961: 6 and 17 Feb (AS.W.. Frost). Hillsborough Co. 112 ) 1964: I-III (A J. Cross). 114 ) 1975: 12 Sept (P). Holmes Co. 116 ) 1975: 28 March (A). 117 ) 1975: 11 June (M,P,C), 6 Sept (S,M,P,C), 30 Dec (S,C). 119 ) 1974: 15 June (S,M,L,P); 1975: 28 March (S) , 11 June (S,M,L,P,C), 6 Sept (S,M,P,C), 30 Dec (M,P,C,A); 1976: 18 April (S,M,L,P,C). Jackson Co. 123) 1939: 9 July (A D.E.Hardy) . Jefferson Co. 124 ) 1975: 26 March (A). 125 ) 1975: 26 March (A), 23 Aug (S).

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-114Lafayette Co. 130 ) 1974: 14 June (P) ; 1975: 26 March (A). Lake Co. 170 ) 21 March (S) , 3 Sept (P) . Leon Co. 131) 1974: 16 March (S,L); 1975: 26 March (L,P,A), 10 June (P,C), 23 Aug (P,C,A), 29 Dec (S) . JL34) 1973: 19 Dec (P) . Levy Co. 135 ) 1973: 7 Oct (P) ; 1974: 2 March (S,M,L), 3 Aug (S,M,L,P, C,A), 15 Sept (S,M); 1975: 31 Jan (S,M), 23 March (P,A), 17 May (P), 30 Oct (S). 136) 1974: 15 Sept (S). 137 ) 1974: 2 March (S), 18 June (S), 3 Aug (S,M), 15 Sept (S,M,L,P); 1975: 23 March (S). 138 ) 1975: 31 Jan (S,M,P,C), 30 Oct (S,A). 139) 1975: 23 March (S.P.C), 11 Sept (S,M,L), 21 Dec (S) . 141 ) 1975: 8 July (P) , 30 Oct (S,M). Liberty Co. 142) 1975: 18 Jan (S,M), 24 Aug (S). 143) 1976: 1-2 April (A-G.B. Fairchild). 147 ) 1975: 24 Aug (M,P). Madison Co. 153 ) 1974: 16 March (S,M,L,P), 5 Aug (S,M,L), 12 Oct (S,M,L,P,A); 1975: 17 Jan (S,M,L), 26 March (S,M,L,P,C,A) , 23 Aug (S,M,L,P,C,A), 29 Dec (S,M). Manatee Co. 156 ) 1975: 12 Sept (S). Nassau Co. 159 ) 1974: 4 Jan (S,P,C), 20 April (S,M,L,P,A), 10 July (S,M,L), 24 Aug (S,M,L,P,C,A), 9 Oct (S,P,C), 4 Dec (C) ; 1975: 12 March (S,M,L,P,A), 4 May (S,M,P,C), 1 July (S,M,L,P) , 26 Sept (S,M,L,P,C); 1976: 14 Feb (S.M.P.A). 160 ) 1974: 20 April (S,P,A), 10 July (S), 24 Aug (S,M), 9 Oct (S,M,L,P,C); 1975: 12 March (S) , 4 May (S.M.P.A), 1 July (C) , 26 Sept (S,M,C). 161 ) 1974: 20 April (M,P,C,A), 10 July (S,M), 24 Aug (S,M,L,P), 9 Oct (S,A); 1975: 12 March (S,M), 4 May (A), 1 July (S) , 26 Sept (S,P); 1976: 14 Feb (C) . 162 ) 1974: 20 April (M,L,P,C), 10 July (L.P.C), 24 Aug (S,M,P), 9 Oct (S,M); 1975: 12 March (S,M,L,P,C), 4 May (A), 22 July (S,M, L,P,C).

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-115Okaloosa Co. 165 ) 1974: 18 March (M,L,P,A), 16 June (S,M,P), 7 Aug (S.M.L.P), 14 Oct (S,M,L,P); 1975: 28 March (S,L), 12 June (S,M). 166) 1974: 18 March (C) , 14 Oct (S) ; 1975: 6 Sept (S). 168 ) 1975: 12 June (S,P), 6 Sept (S,C). Orange Co. 169 ) 1973: 11 Sept (S,P); 1974: 13 July (S,M,L,P); 1975: 15 March (P) , 4 July (S,P), 30 Oct (S,M,L,P,C). 174 ) 1936: 31 March (L,P U. S.N.M. ) ; 1941: 13 March (L, P, A W.V.King) ; 1947: 28 Jan (P,A H.K. Gouck) , 5 and 11 Feb (P,A H.K. Gouck) , 10 March (A H.K. Gouck), 13 March (P H.K. Couck) ; 1953: 5 Aug (A K.M. Sommerman), 4 Oct (AK.M. Sommerman) . 176 ) 1936: 31 March (A). Pasco Co. 177) 1975: 23 March (P,C,A), 21 Dec (S,M,L,P,C,A) . 178 ) 1939: 13 July (A D.E. Hardy). Osceola Co. 179) 1932: 1 Feb (A A.L. Melander) . Pinelas Co. 180 ) 1936: 5 March (A — collector unknown). Polk Co. 182) 1975: 27 May (S,M). Putnam Co. 185 ) 1973: 17 Nov (S,M,L,P,C,A) ; 1974: 5 Jan (S,M,L,P,A), 14 Feb (S,M,L,P), 26 March (S,M,L,P,C), 18 May (S,M,L,P,C,A) , 6 July (S,M,L,P,A), 12 Aug (S,M,L,P,C,A), 6 Oct (S,M,L,P,A), 23 Nov (S,M, L,P,C); 1975: 15 Jan (S,M,L,P,C,A) , 4 April (S,M,L,P,C), 11 May (S,M,L,P,C), 14 June (S,M,L,P), 17 July (S,M,L,P,C,A) , 30 Sept (S,M,L,P,C,A), 14 Dec (S,M,L,P,C,A) . 186 ) 1974: 19 Jan (S,M,L,P,A), 14 April (S,M,L,P,C), 25 May (S,M,L,P), 6 July (S,M,L,P,C,A) , 17 Aug (S,M,L,P,C), 6 Oct (S,M,L,P,C,A), 23 Nov (S,M,L,P,C); 1975: 12 Feb (S,M,P,A), 18 April (S,M,P), 26 June (P,C), 25 Sept (S,M,L, P,C). 187 ) 1974: 6 July (P) . 188 ) 1964: 9 April (A H.A. Denmark). 190 ) 1974: 7 Dec (S,M,C). 192 ) 1974: 6 Oct (S) , 7 Aug (C) . 193) 1974: 26 Jan (S,M,L,P,C,A) .

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-116Santa Rosa Co. 195 ) 1975: 12 June (P) . 196 ) 1974: 16 June (S) ; 1975: 12 June (S). 197) 1973: 23 May (A W.W. Wirth) . Sarasota Co. 198 ) 1967: 13 March (S,L W. Beck). Seminole Co. 203 ) 1974: 21 March (S,M,L,P), 12 May (S,M,P), 11 July (M,L,P), 3 Sept (S,M,L,P,C), 28 Nov (S,M); 1975: 15 March (L,P), 4 July (S,M,L,P,C), 31 Oct (S,M,L,P,C). 204 ) 1960: 15 and 31 March (Larvae — E.G. Jay). Sumter Co. 206) 1973: 21 Nov (M,L,C); 1974: 28 Nov (S,M,P). Suwanee Co. 209 ) 1945: 10 Jan (A D.J. Taylor). Taylor Co. 210 ) 1954: 8 April (P CM. Jones); 1974: 16 March (S,M, L), 14 June (S,M,L,P), 5 Aug (S.M.L.P), 12 Oct (S,M,L,P); 1975: 17 Jan (S,M,L,P), 26 March (S,M,L,P), 10 June (S,M,L,P,C), 23 Aug (S,M,L,P,C). 211 ) 1974: 16 March (S,L,P), 14 June (S,M,L,P), 5 Aug (S,M,P), 12 Oct (S,M,L,P); 1975: 17 Jan (S,M,L,P,C), 26 March (S,M,L,P,C,A) , 10 June (S,M,L,P,C), 23 Aug (S,M,L,P,C), 29 Dec (P,C). 213 ) 1974: 14 June (S,M,L,P), 5 Aug (M,P) , 12 Oct (P) ; 1975: 17 Jan (S,L), 23 Aug (S,P). 214 ) 1975: 17 Jan (S,M,L). 215) 1975: 26 March (S,L,P,C,A), 23 Aug (S,M,L,P,C). Union Co. 216 ) 1974: 5 Jan (S), 23 Feb (S,M,L,P,C), 14 April (S,M,L,P,C), 1 June (S,M,L,P), 6 July (S,M,L,P), 24 Aug (S,M,L,P,C), 9 Oct (S,M,L,P,C), 6 Nov (S,M,L,P,C,A); 1975: 12 Jan (S,M), 5 April (S,M,L,P,C,A), 4 May (S,M,L,P,C,A) , 6 May (A), 11 May (A), I June (S,M,L,P,C), 15 June (S,M,L,P,A), 22 July (S,M,L,P,C), 5 Oct (S,M, L,P,C), 14 Nov (S,M,L,P,C); 1976: 7 Jan (S,M,L,P,C,A) , 17 Feb (S,M,L,P,C,A), 24 March (P,A). 217) 1974: 23 Feb (A), 6 July (S,P,C), 24 Aug (S.L.P.C), 9 Oct (S,M,P,C); 1975: 1 June (S,L,P,C), 22 July (P), 5 Oct (S,M).

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-117Wakulla Co. 219) 1970: 29 April (A W.W. Wirth). Walton Co. 220 ) 1974: 13 Oct (L) ; 1975: 11 June (P,A). 221) 1975: 11 June (M), 6 Sept (M) . 222 ) 1974: 6 Aug (S,M,A). 223 ) 1975: 28 March (S), 11 June (S,L). Simulium (Eusimulium) aongareenarum (Dyar and Shannon) Eusimuliwn aongareenarum Dyar and Shannon, 1927, Proc. U.S. Nat. Mus. 69(10): 20 (female). Simulium aongareenavum — Jamnback and Stone, 1957, Ann. Entomol. Soc. Amer. 50: 395 (male, female, larva, pupa). Simulium congareenarum — Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 27 (male, female, larva, pupa). Taxonomy . Dyar and Shannon (1927) first described a female of S. aongareenavum (as Eusimulium aongareenarum) and the holotype location was given as Congaree, South Carolina. The holotype and twenty-three paratype females were deposited in the U.S. National Museum. Jamnback and Stone (1957) provided the first descriptions for the larva, pupa, and male of S. congareenarum and redescribed the female. Davies et al. (1962) mention that morphological and cytological evidence suggests that S. aongareenavum is a species complex. Stone and Snoddy (1969) indicate that biological information on hand also suggests a complex exists. Description . Mature larvae are 5.5-6 mm long with a yellow head capsule and dark brown head spots. The anterior medial group of spots is made up of 4 or 5 small distinct spots which are widely separated from the posterior medial group of spots (Fig. 33) . Each cephalic fan

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-118.1 > Figure 33. Cephalic apotome of a 5. congareenavum larva. Figure 34. Venter of the larval head capsule of S. congareenamtm .

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-119contains 57-61 rays. The gular notch is in the form of a small square or is of a shallow U-shape and extends less than one-fifth the distance to the submental teeth (Fig. 34). The abdomen is reddish brown with lighter intersegmental areas. The ventral, anal tubercles are large and cone-shaped. The pupa is about 3 mm long and bears a pair of 12-f ilamented respiratory organs (Fig. 35). The anterior edges of the cocoon in a lateral view are concave and a long anterior projection occurs dorsally on the cocoon. The wing of the male is 2.5 mm long. The scutum of the male is velvety black but is covered with many golden hairs. The scutellum bears long erect silvery or golden hairs. The base of the radius bears hairs dorsally. The ventral plate in end view is a good sized sharppointed V. The distimeres are only slightly curved and taper to a sharp point (Fig. 36). The wing of the female is about 2 . 5 mm long. The frons and clypeus are gray pollinose in appearance. The scutum appears gray with many closely appressed pale hairs. The wings bear setae on the base of the radius and there is a small basal cell. The pedisulcus on the second tarsal segment is shallow. The tarsal claw has a prominent basal tooth with convex margins. The genital fork has stout arms and prominent inner projections (Fig. 37). Distribution . In addition to its occurrence in South Carolina 5. oongareenarum has been reported from New York, Florida (Alachua Co.), Georgia, Louisiana, Maryland, and Virginia (Jamnback and Stone, 1957), Ontario, Canada (Davies et al. , 1962), Connecticut (Stone, 1964), Alabama (Stone and Snoddy, 1969) and New Jersey (Crans and McCuiston, 1970a).

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-120Figure 35. Pupa and cocoon of S. eongareenamm. Figure 36. Terminalia of a male of S. eongareenamm.

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-121--*, , V #fr Figure 37. Terminalia of a female of 5. congareenarum.

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-122Lifo history . In Canada overwintering eggs of the first generation begin hatching in early April, pupae and adults are present by May and a possible second generation is indicated by a peak of pupation during July (Davies et al. , 1962). Stone (1964) reports that adults from the overwintering larvae emerge during early March in the southern regions and in April in the north. In South Carolina larvae of S. congaveenavum were collected during January, were abundant in February and March and adults were present during March through early May (Jones and Richey 1956; Anthony and Jones, 1958; Noblet et al. , 1972). Jones and Richey (1956) also report collecting a few larvae and pupae through 11 June and one larva on 13 July. In South Carolina 5. congaveenavum appears to be a univoltine species (Garris et al., 1975). A 20to 25-day life cycle from egg to adult emergence is reported for S. congareenavwn and other Si.nm.lium species near Pageland, South Carolina (Kissam et al., 1975). Ecolo gy. Jamnback and Stone (1957) collected S. congareenavwn in a permanent creek .61 m (2 ft) wide and 15 cm (6 in) deep. Stone (1964) indicates that the immatures of S. .congaveenavum attach to vegetation in fairly slow-flowing permanent streams. Garris et al. (1975) indicate that this species usually occurs in swampy streams. Habits. During early April S. congaveenavwn adults were collected from breeder turkeys (Anthony and Richey, 1958). The number of adults attracted to turkeys decreased to zero by mid-May (Jones and Richey, 1956). Noblet et al. (1972) found that the adults feed on turkeys, ducks, and chickens and showed, by injection of homogenized infected flies, that S. congaveenavwn was a vector of L, smithi to turkeys. Noblet et al. (1975) report that 8. congaveenavum is an important vector of L. smithi in the early spring.

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-123Florlda observations . Stream Parameters Width Depth pH Temperature VelocityMean: 2.55m 30.97 cm 4.3 ' 16°C (61°F) .46 m/sec (1.51 ft/sec) Min: .076 1.27 3.5 6.7 (44) .15 ( .5) Max: 11 166 6.7 26.1 (79) 1 (3.3) Figure 38 shows the distribution of S. congareenarum in Florida where it was found in 16 counties at 34 sites. This species was only collected in the northern portion of the state. One young larva was collected on 6 September at Little Reedy Creek (Site 117) in west Florida and a large larva was recovered from Water Oak Creek (44) in east Florida on 26 September. At all other locations favorable for S. congareenarum larvae were not found in the fall before early October. Larvae and pupae were present in many streams in greatest numbers from January through March and occasionally into April. By May the numbers of S. congareenarum were significantly lower, both in permanent streams where S. slossonae also occurred and was building up its populations, and in temporary streams where the flow reached its lowest level or ceased prior to the summer rainy season. The latest record for larvae was 11 June at Site 117. Adults were captured during January, March, April and late May. It appears that S. congareenarum is able to complete at least two and possibly three generations each breeding season and usually spends the summer in the egg stage or possibly, in more permanent flows, as small numbers of larvae. The streams in which the largest populations of S. congareenarum were encountered (8, 43, 44, 117, 131, 153, 211, 216 Fig. 39) were usually 1 to 3 in wide 10 to 50 cm deep with currents .46 to .76 m/sec

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-124Figure 38. Collection locations for S. aongareenarum in Florida.

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-125•* -t^ . P ft-* n, ; • V .'':: , W ^mST'Si Figure 39. Site 216, Turkey Creek, a typical 5. aongareenarum stream.

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-126(1.5-2.5 ft/sec) and a low pH, 3.5-4.5. These streams had trailing grass as a primary type of vegetation in the flow and, generally, sandy bottoms. Larvae and pupae were found attached to grass, sedges, large and small-leafed aquatic vegetation, dead leaves, pine needles and, occasionally, rocks. Simuliwi congareenarum was collected with nine other black fly species (Table 3 ). Most notably it occurred with S. slossonae and S. tuberosum and was recovered frequently with two other winter and early spring species, C. ornithophilia and S. verecundum. Infected larvae with tiny white spheres in the abdomen and thorax or a large white mass in the abdomen were noted only a few times in collections of S. congareenarum and only involved 1-3 larvae per collection. Adults reared in the laboratory during April 1975 and March April 1976 which were dissected before taking a blood meal were found to have well-developed eggs in the ovaries. This suggests some females of S. oonaareenarum may be autogenous at least for the first ovarian cycle. It may also help explain the reluctance of reared S. congareenarum to feed on turkeys in the laboratory while wild-caught S. congareenarum, which may have already oviposited, fed more readily. Adults fed a number of times on turkeys in the field and more often in the laboratory and transmitted L. smithi to clean birds on three occasions. This is the second most important vector of L. smithi to turkeys in Florida. The vector potential of S. congareenarum is limited by its seasonality and the relatively lower number of adults in flight. Adults were captured in Manitoba traps during January-March at four sites in four counties (Table A), in a Malaise trap at Site 2 in March,

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-127and from exposed turkeys on a few occasions during early spring (Table 8 ). Simulium oongareenarvm was easy to differentiate in the field from the frequently collected S. slossonae by the more robust appearance of the former, its more awkward ambling up a Manitoba canopy, and its silvery gray frons. Florida collection records for S. aongareenarum . Alachua Co. 1) 1973: 7 Dec (L) , 13 Dec (L) ; 1975: 12 Feb (S) ; 1976: 21 Jan (S). 2) 1975: 19-20 March (A-G.B. Fairchild) . 6) 1974: 2 April (S); 1975: 8 Oct (M) . 8) 1974: 14 Feb (S) , 2 April (S), 16 May (S); 1975: 10 Jan (S,M), 12 Feb (M,P,C,A), 9 Nov (S) , 14 Dec (S); 1976: 6 March (M) . 14) 1974: 7 March (S) , 30 May (S,L). 2J.) 1974: 2 March (S) , 11 April (S,L); 1975: 4 April (M,C); 1976: 21 Jan (S,M), 29 Jan (S,P,A). 22) 1975: 24 Jan (A); 1976: 6 March (A). 23) 1975: 24 Jan (S,M). 36) 1975: 15 Jan (L) , 4 April (M) , 22 Oct (S). 38) 1957: (no month or stage — Jamnback and Stone). Baker Co. 41) 1974: 4 Jan (S); 1975: 1 Feb (S) . Bradford Co. 42) 1974: 4 Jan (S) . 43) 1973: 17 Nov (S,M,P,A); 1974: 4 Jan (S,M), 16 Nov (P,A); 1975: 12 Jan (S,M,L,P,C), 26 Jan (A), 5 April (S,M,L,C); 1976: 7 Jan (S,M,P,C,A), 31 Jan (S,M,L). 44) 1974: 4 Jan (S), 23 Feb (C), 14 April (S,M,L); 1975: 12 Jan (S,L), 5 April (S) , 26 Sept (L) ; 1976: 31 Jan (S,M). Duval Co. 2°) 1974: 4 Jan (S,M,P); 1975: 12 March (S) . Franklin Co. 80) 1976: 2-3 April (A-G.B. Fairchild). Hamilton Co. 99) 1976: 14 Feb (M) . Holmes Co. 117 ) 1975: 19 Jan (S,M,L,P), 28 March (S,M,L,P), 11 June (S,M), 6 Sept (M), 30 Dec (S,M). 119 ) 1975: 19 Jan (S,M,L,P), 28 March (S,M,L,P,A), 30 Dec (S,M,L,P,C); 1976: 18 April (S) .

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-128Lafayette Co. 130 ) 1975: 17 Jan (S), 26 March (C) . Leon Co. 131 ) 1973: 17 Dec (S,M,L); 1974: 16 March (S,M,L,P,C); 1975: 26 March (S,L), 29 Dec (S,M,L). Liberty Co. 143) 1976: 1-2 April (A-G.B. Fairchild) . 144 ) 1973: 18 Dec (S). 147 1974: 17 March (S) ; 1975: 18 Jan (S,M), 27 March (S,M), 30 Dec (P,C,A). Madison Co. 153 ) 1974: 16 March (S,M,L,P,C), 12 Oct (M) ; 1975: 17 Jan (S,M,L,P,C), 26 March (S ,M,L,P,C,A) , 29 Dec (S,M,L). Nassau Co. 159 ) 1974: 4 Jan (S); 1975: 12 March (A); 1976: 14 Feb (S,M,L). 160 ) 1974: 4 Jan (S), 20 April (S,M,L); 1975: 12 March (S,M). 161 ) 1975: 12 March (S,M); 1976: 14 Feb (L) . 162) 1975: 12 March (P,A). Okaloosa Co. 168 ) 1973: 13 March (S,M,L,P K. Tennessen) . Santa Rosa Co. 197 ) 1973: 23 May (A W.W. Wirth) . Taylor Co. 210 ) 1974: 16 March (M) ; 1975: 17 Jan (S,M,L), 26 March (S,M,L,P), 28 Dec (S,M). 211 ) 1974: 16 March (S,M,L,P); 1975: 17 Jan (S,M,L), 26 March (S,M,L,P), 29 Dec (P) . 214 ) 1975: 17 Jan (S,M,L,P). 215 ) 1975: 26 March (S,M,L). Union Co. 216 ) 1974: 5 Jan (S,M,P), 23 Feb (S,M,L,P,C), 14 April (S,M,L,P), 9 Oct (S), 6 Nov (S,M); 1975: 12 Jan (S,M,L,P,A), 5 April (S,M,L,P,A), 4 May (S,M,C), 14 Nov (S); 1976: 7 Jan (S,M, L,P,A), 17 Feb (S,M,L,P,C,A) , 24 March (P,A). Simulium (Phostevodovos) dixiense Stone and Snoddy Simulium dixiense Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 34 (female, male, pupa).

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-129Taxonomy . A female of S. dixiense , with the associated pupal exuvium and cocoon, was designated as the holotype (Stone and Snoddy, 1969). The type locality is Lewis Creek, Washington Co., Alabama. The holotype is in the U.S. National Museum. Description . Stone and Snoddy (1969) indicated that unless the respiratory filaments of well-developed larvae were examined, they were probably not distinguishable from S. jonesi or S. nyssa. During this research S. dixiense was found to be readily distinguished from other species and had the following characteristics: the larva is 4.5 mm long; the color of the larval head capsule and abdomen is a striking light yellow with the larvae appearing almost clear and very difficult to detect on eel grass except for the dark histoblasts of well-developed larvae and the sclerotized anal X-piece and the mandibles; the head capsule is conspicuously elongate; the head spots are fairly faint and light brown in color and near pupation these spots disappear and are replaced by whitish yellow or colorless areas (Fig. 40); the gular notch is elongate, extends two-thirds the distance to the submental teeth, is widest about halfway along its length but not nearly as wide as long, and is pointed anteriorly (Fig. 41); the hypostomium bears a dark sclerotized band at the base of the teeth; the cephalic fans consist of 55-58 rays each; the shape of the fully open fan is more like an elongate kidney bean than a semi-circle; the antennae extend slightly beyond the cephalic fan stalk; the abdomen is swollen and curved posteriorly giving the anal hooks a more ventral than terminal placement; there are approximately 63 rows of anal hooks with 12 hooks in each row; the anal tubercles are prominent; the anterior arms of the anal X-piece appear long; and the body in alcohol is bowed or curved into a C-shape (Fig. 11).

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-130Figure 40. Dorsal view of the head capsule of a S. dixiense larva. Figure 41. Gular notch and hypostomium of a S. dixiense larva.

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-131The pupa Is 2.5-3 ram long with filaments 1 mm long. The respiratory organ consists of 10 filaments in 5 pairs. The fourth petiole from the dorsal is the thickest and longest extending 1/3 or more of the length of the filaments. The anterior margins of the cocoon in lateral view appear straight, not convex, and slope back from the ventral to the dorsal. There is a large aperture on each side of the cocoon anteriorly (Fig. 12). Pupae often appear bright yellow when freshly collected. The male bears wings 2-2.5 mm long. The scutum has a pair of large silvery spots anterior laterally and a wide, posterior silvery area with the remainder of the scutum being velvety black and expanding anteriorly between the two spots. The abdomen is velvety brownish black with iridescent spots along the side. According to Stone and Snoddy (1969) the terminalia are not distinguishable from those of S. jenningsi (Fig. 42). The female's wings are 2-2.5 mm long. The stem vein hairs are dark and there are no setae on the base of the radius. The frons is shiny dark brown. The scape and pedicel of the antennae are orangishred and the flagellum is dark brown. The scutum is shiny dark brown with many small golden hairs and appears poilinose laterally. The legs are dark distally, lighter yellow or yellow brown proximally. The abdomen is velvety black except the last four tergites which are shiny dark brown. The stem of the genital fork is thin and dark while the arms are wider, lightly sclerotized and end in a dark sclerotized tip (Fig. 43). Distribution . Stone and Snoddy (1969) report S. dixiense from Alabama and South Carolina. Life history . Stone and Snoddy (1969) suggest this species is multivoltine and report collecting pupal exuviae in early March and

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-132Figure 42. Male terminalia of S. dixiense. Figure 43. Terminalia of a female of 5. dixiense.

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-133larvae and pupae into November. Garris et al. (1975) in a survey conducted from May through October in Sumter County, South Carolina, collected pupae of S. dixiense only in May and July. Ecology . Stone and Snoddy (1969) found that S. dixiense prefers the small swamp streams of the Southeast and collected this species in Alabama only from the Lower Coastal Plain. Larvae and pupae were usually collected in sand-bottomed streams 15-20 cm (6-8 in) deep and .92-1.2 m (3-4 ft) wide and seemed to prefer grasses, Zvcnaus sp. and other green vegetation for attachment sites especially in stream sections where a constriction increased the velocity to .31-. 92 m/sec (1-3 ft/sec) (Stone and Snoddy, 1969). Stone and Snoddy (1969) report that S. dixiense is more commonly collected with S. slossonae than S. jonesi, apparently preferring smaller cooler streams than S. jonesi prefers. Tarshis and Adkins (1971) report successfully transporting S. dixiense and other black fly larvae in aerated water in coffee cans in a styrofoam chest for distances up to 200 miles. Habits . Nothing is reported in the literature on the adult habits. Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 7.1 m 52 cm 4.25 19°C (66.2°F) .67 m/sec (2.2 ft/sec) Min: .6 8 3.75 13.6 (56.5) .31 (1) Max: 15.3 100 + 4.55 25 (77) 1.22 (4) Simulium dixiense was collected from seven locations in four west Florida counties (Fig. 44). The solid dots indicate sites where welldeveloped, large larvae and/or pupae of S. dixiense were found. The open circles designate collection sites where small larvae displaying

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-134r/ J&* Figure 44. Collection locations for S. dixiense in Florida.

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-135Figure 45. Site 74, Pine Barrens Creek, a stream inhabited by 5. dixiense.

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-136S. dixiense characters were recovered and the presence of this species should be further supported by collections of later immature stages. Large larvae and pupae have been collected from January through December indicating year round breeding for this species and multiple generations. Simulium dixiense was typically found in streams 7-15 m wide and .5 m to 1.0 m deep or deeper in sections. Immatures were recovered from the surface of the flow to about 70 cm deep. The streams exhibited a low pH reaction usually 4.0-4.5 and immatures were only collected in temperatures from 13.6°C (56.6°F) to 25°C (77°F). The stream velocity was moderately swift, usually about .46-1 m/sec (1.5-3.3 ft/sec). A typical collection site is Pine Barrens Creek (Site 74) in Escambia County (Fig. 45). Larvae and pupae were most commonly found on clean eel grass but also were collected from eel grass covered with a thin brown sediment layer or a gelatinous layer of green algae. Immatures were also found attached to thin and wide trailing bank grass, dead tree leaves, small sedges, and long thin green reeds, triangular in cross section. Simulium dixiense was associated with five other black fly species: most frequently with S. jonesi but also with S. tuberosum, S. vereoundum, S. stossonae, and S. haysi. Mermithid nematodes were observed on two occasions during October in three small larvae and during late December in two small larvae, in all cases at Pine Barrens Creek, Site 74. Florida collection records for S. dixiense . Calhoun Co. _49) 1975: 27 March (S,M,L,P), 6 Sept (P) . Escambia Co. 13) 1974: 18 March (S,M,P), 16 June (P) . 74) 1974: 16 June (S,M,L,P), 7 Aug (S,M,P,C), 14 Oct (S,M,L,P); 1975: 19 Jan (S,L,P,A), 28 March (S,M,L,P,C,A) , 12 June (S,M,L,P,C,A) , 7 Sept

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-137(S,M,L,P,C,A), 31 Dec (S,M,L); 1976: 18 April (S,M,L,P,A). Okaloosa Co. 166 ) 1975: 28 March (S), 6 Sept (S), 31 Dec (S). 168) 1975: 29 March (S) . Santa Rosa Co. 195) 1974: 18 March (S,M), 7 Aug (S,M) , 14 Oct (L,P) ; 1975: 28 March (S.M.P.A), 7 Sept (P,C), 31 Dec (P). 196) 1974: 18 March (S) ; 1975: 31 Dec (S). Sinruliiim (Phos te rcdoros) hay si Stone and Snoddy Simuliwn hay si Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 36 (female, male, pupa, larva). Taxonomy . The holotype of this species is a female from Burnt Corn Creek, Brewton, Escambia Co., Alabama. The holotype female with pupal exuvium is deposited in the U.S. National Museum (Stone and Snoddy, 1969). Description. The larva is 4.1 mm long. The head capsule and abdomen in a preserved state are light brown in color. The head spots are slightly darker brown and typically arranged (Fig. 46). The gular notch is broad, widest about midway along its length, extends just over half the distance to the teeth of the submentum and is pointed apically (Fig. 47). The antennae extend beyond the length of the fan stalks. The anal gills are arborescent. The ventral tubercles are large. The pupa is 2.5-3 mm long. The head bears at least two large simple trichomes. The respiratory organ consists of 7 filaments, 6 of which arise in pairs from the basal 7th filament (Fig. 48). The wings of the male arc 1.75-2 mm long. The male is black in appearance with two silvery, iridescent anterior spots on the scutum

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-133Figure 46. Cephalic apotome of a S. haysi larva. Figure 47. Gular notch of a S. haysi larva.

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-139Figure 48. Pupal exuvium and cocoon of S. haysi.

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-140which diverge anteriorly and posteriorly. The abdomen is black with silvery patches on the side. Stone and Snoddy (1969) indicate the tenninalia are as in S. jonesi. The female is small and black in appearance. The wings are almost 2.0 mm long. The frons in shining black. The clypeus is longer than wide. The scutum is dark, shiny black without pollinose anterior spots. The tibia bear shiny white patches. The genital fork has a thin stem and thin arms which branch out almost perpendicular from the stem. Distributio n. This species has been reported from a few locations in Alabama (Stone and Snoddy, 1969). Life history . In Alabama S. haysi overwinters in the egg stage, is common from June to September, and completes three or more generations per year (Stone and Snoddy, 1969). Ecology . Immatures of S. haysi have been collected from sedges and grasses in swift flowing, shallow sections of large streams of the Lower Coastal Plain of Alabama. Habits . Nothing is reported on the habits of the adults in the literature. Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 12.4 m 52.8 cm 4.1 21.1°C (70.1°F) 1 m/sec (3.3 ft/sec) Min: 5 15 3.75 17.2 (63) .436 (1.43) Max: 15.25 100 4.4 22.8 (73) 1.38 (4.54) Mature larvae and pupae of S. haysi were recovered from only one stream in Florida, Big Juniper Creek, Site 195 (Fig. 49). The stream parameters above summarize stream conditions prevalent when the immatures were

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-141r, y JiT» Figure 49. Location of the collection site for S. haysi in Florida.

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-142Figure 50. Site 195, Juniper Creek, where S. haysi was collected.

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-143collected at this site. The earliest collection occurred on 12 June and the latest on 14 October. Big Juniper Creek (Fig. 50) is a fairly large, rapidly flowing stream with a firm yellow clay or limestone substrate and large rock-like clay boulders often covered with a thick layer of slippery green algae. The creek is located in a steep-sided tree and bush-covered gorge about 20 m below the highway. The flow is shallow on the east side but on the west side a channel 1 m deep occurs. Little green vegetation occurs in the flow except some small trailing tree and bush leaves and a few sedges and clumps of grass. Immatures were found on the hard, rock-like clay and on twigs. Only small numbers of mature larvae and pupae, never more than twenty specimens per visit, were collected although some of the small unidentifiable Phosterodoros larvae collected at the same times may include more representatives of this species. Simulium haysi was associated with S. jonesi, S. tuberosum, S. dixiense and S. slossonae at this location. At the Chipola River, Site 50, in Calhoun County, one medium-aged larvae with white histoblasts was collected in March. When mounted on a slide these histoblasts appeared to have seven filaments like the respiratory organ of S. haysi. Three subsequent visits to this site yielded no further S. haysi-llke specimens. Florida collection records for S. haysi . Santa Rosa Co. 195 ) 1974: 16 June (L,P), 7 Aug (L,P,A), 14 Oct (L,P,A) 1975: 12 June (P) , 7 Sept (P) . Simulium (Phosterodoros) Jenningsi Malloch Simulium jenningsi Malloch, 1914, U.S. Dept. Agr. Bur. Entomol. Tech. Ser. 26: 41 (female, male, larva, pupa).

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-144Simulium nigroparvum Twinn, 1936, Can. J. Res., D, 14: 142 (female, male, pupa). Simulium nigroparvum — Underhill, 1944, Va. Agr. Exp. Sta. Tech. Bull. 94: 1-32 (female, male, pupa, larva). Simulium jenningsi — Davies, Peterson, and Wood, 1962, Proc. Entomol. Soc. Ontario 92: 118 (female, male, pupa). Simulium jenningsi — Wood, Peterson, Davies, and Gyorkos, 1962, Proc. Entomol. Soc. Ontario 93: 112 (larva). Simulium jenningsi — Stone, 1964, Conn. State Geol. and Natur. Hist. Surv. Bull. 97: 44 (female, male, larva, pupa). Taxonomy . Malloch (1914) described a female of S. jenningsi as the holotype which was deposited in the U.S. National Museum. The type locality is Plummers Island, Maryland (Davies et al. , 1962). Stone and Snoddy (1969) created the subgenus Phosterodoros to encompass the species in the former S. jenningsi complex which are basically similar and likely share a common ancestor, possibly the tolerant and widespread S. jenningsi. Description . The larvae are about 5 mm long. The abdomen is light brown, sometimes with a reddish hue. The head spots are dark on a light yellow brown head capsule (Fig. 51). The throat cleft is broadly sagittiform, usually pointed anteriorly and extends about halfway to the submental teeth (Fig. 52). The respiratory histoblasts contain ten thin filaments. The pupa is about 2.5 mm long. The respiratory organs each consist of ten filaments in a 2, 2, 3, 3 arrangement from dorsal to ventral. The cocoon is tightly woven, slipper-shaped and has a large aperture on each side anteriorly (Fig. 53).

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-145=jfo«-»* " Figure 51. Cephalic apotome of a 5. jennzngsi. larva. Figure 52. Gular notch of a S. jenningsi larva.

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-146Figure 53. A pupa and cocoon of S. jenningsi. Figure 54. S. jenningsi male terminalia.

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-147| V I Figure 55. Genitalia of a female of S. jenningsi.

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-148The male, about 2.25 mm long, is velvety black in appearance with shiny silvery areas at both the anterior lateral angles and on the posterior third of the scutum. The ventral plate in ventral view is longer than wide, in end view displays a median section roundly elongate and knobbed or pointed distally; the distimeres are three times longer than wide and tapering (Fig. 54). The female tfing is about 2.1 mm long. The female has a shiny dark brown scutum with many tiny golden appressed hairs and gray pollinosity especially laterally. The frons is shiny dark brown. The scutellum bears many long erect dark hairs posteriorly. The base of the radius and underside of the subcosta are bare. The fore coxa and femur are light yellow contrasting with the dark brown tarsal segments and distal tip of the tibia. The arms of the genital fork are fairly thin, widely separated with a slender dark dorsal projection and a more broad weakly sclerotized ventral projection (Fig. 55). Distribution . Stone (1964) reported that S. jenningsi was widely distributed in eastern North America from Manitoba to Maine and south to Texas and Florida. Stone and Snoddy (1969), mentioning difficulties in separating adults in the subgenus Phosterodoros , restricted the distribution of S. jenningsi to pupal records and listed the species from Alabama, Connecticut, Kentucky, Maryland, Michigan, New York, Virginia, Wisconsin and Ontario, Canada. Crans and McCuiston (1970a) report S. jenningsi from New Jersey. Life history . Underhill (1939) collected immature stages of 5. jenningsi (as S. nigroparvian) in streams from April until November and suggested overwintering occurred as an egg or immature larva. Stone and Snoddy (1969) found no larvae during late winter and suggested that eggs

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-149overwinter with hatching occurring in March or April. Immature stages of S. jenningsi were found in central Alabama from June through November (Stone and Snoddy, 1969). Anderson and Dicke (1960) report that in the cooler water the first generation larvae require five to six weeks to mature, indicate the pupae take five to seven days to develop and mention that the adults of the first generation in Wisconsin are usually on the wing by the third week in May. Stone and Snoddy (1969) report that five or more generations are completed each year in Alabama. Ecology . Underhill (1944) found the immature stages of S. jenningsi (as S. nigroparV'urn) to be most abundant in clear streams at least 7.69.2 m (25 to 30 ft) wide, 28-61 cm (15-24 in) deep that flowed 1.2-1.8 m/sec (4-6 ft/sec) and contained trailing water willow Dianthera (now Justicia) amerioana. Anderson and Dicke (1960) found immatures in the clear, shallow, rocky, swift portions of young rivers 2.5-15 cm (1-6 in) below the surface on vegetation and 46 cm (18 in) deep on rocks. Stone and Snoddy (1969) mention S. jenningsi prefers the rapids sections of unpolluted large inland streams and rivers and immatures are found in water about 15-28°C and nearly neutral in pH. Habits . Underhill (1944) suggested that adults may travel 32-48 km (2030 mi) from their breeding locations. After being unable to find egg masses Underhill (1944) proposed that females deposited eggs into the streams while flying. This species has been observed in Virginia feeding on turkeys, horses, mules, cattle and, rarely, on man (Underhill, 1939, 1944). Stone (1964) mentions that it is difficult to separate 5. jenningsi from closely related adults and suggests records of feedings on turkeys may actually be the activities of a close relative. The favorite hosts are probably cattle and horses (Stone, 1964). Stone and

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-150Snoddy (1969) report that S. jenningsi is an annoying pest in the vicinity of Washington D.C. throughout the summer. Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 9. 42 m 48.93 cm 6.05 21.3°C (70.4°F) .52 m/sec (1.71 ft/sec) Min: .25 2 3.75 9.4 (49) .15 ( .5) Max: 100 500 7.55 29.4 (85) 1.78 (5.85) Simulium jenningsi is reported from 21 counties and 33 different locations in Florida at all times of the year with records based primarily on pupae and mature larvae with characteristic respiratory histoblasts (Fig. 56). Site 63 is marked at Biscayne Bay in Dade Co. to call attention to a record in Malloch (1914) of an adult, apparently of this species, collected by Mrs. A.T. Slosson. Simulium jenningsi sometimes occurs in fairly wide or very wide and deep flows such as the Santa Fe River (Site 58), the Alapaha River (99), the Waccasassa River (136, 137) and the Withlacoochee (151) but often just sporadically or consistently in low numbers. Larger and more constant populations were encountered in smaller streams, generally under 5 m wide, with a fairly wide range of current characteristics: a flow to Newnan's Lake (18), Joshua Creek (65), Banana Creek (106), Blackwater Creek (113), Kettle Creek (130), Site 141 at Gulf Hammock (Fig. 57), Howell Creek (202) and Rocky Creek (213) . These streams exhibited a pH approaching neutrality (about 6.5-7.5) and all contained some trailing vegetation, often grass. In other respects such as substrate (sand-mud, sand, shells, rocks, concrete) and velocity (.3-1.83 m/sec = 1-6 ft/sec) they varied considerably. A tendency for larger populations to occur

PAGE 164

-151r/ J0*> Figure 56. Collection locations for S. jenningsi in Florida.

PAGE 165

-152Figure 57. Site 141 at Gulf Hammock where S. jenningsi was collected.

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-153in faster flows, .53-. 92 m/sec (1.75-3 ft/sec) was observed. Immaturcs were found attached to grass, eel grass, cattail blades, trailing water willow leaves, a wide variety of additional aquatic vegetation including trapped water hyacinths plus dead leaves, pine needles, shells, and rocks. S. jenningsi was associated with 10 other black fly species, most frequently with S. lakci and S. taxodium and much less frequently with S. slossonaCy S. tuberosum and S. jonesi. This species has multiple generations each year in Florida. It was noted that with the Florida S. jenningsi pupae the 5th (from the dorsal) respiratory filament consistently rises dorsally from the common base of filaments 6 and 7 while figures in the literature (Underhill, 1944 and Stone and Snoddy, 1969) illustrate a ventral origin for the single filament of the first triplet. The single filament of the 2nd (most ventral) triplet rises ventrally as typically illustrated. The surface of the respiratory filaments appears to be more smooth than the rather sharply ridged surface described for S. jenningsi in other areas and the filaments appear speckled like those of S. nyssa. Variations in the number of filaments from the normal 10 and 10 (9 and 10, 10 and 11) have been noted as has considerable difference in the point of bifurcation of the filaments. The ventral plate (Fig. 54) appears more like that illustrated for S. jenningsi in Davies et al . (1962) than in Stone and Snoddy (1969). Florida co 1 lection records for S. jenni.rigsi. Alachua Co. 18) 1974: 12 Jan (P) , 30 June (P) , 24 July (P) , 24 Aug (P) , 22 Sept (P), 25 Oct (L,P), 23 Nov (P) ; 1975: 11 May (P) , 14 June (L,P), 1 Sept (L), 17 Oct (P) , 10 Doc (P) . 20) 1974: 6 July (P) , 29 Aug (P); 1975: 31 Jan (P) , 18 April (L,P), 26 July (P) , 3 Oct

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-154(L,P,A). 21) 1974: 11 April (P) ; 1975: 24 Jan (L) , 27 Sept (P) . 22) 1974: 12 Sept (P) ; 1975: 29 April (P), 30 Aug (P) , 17 Oct (P). 24) 1974: 25 May (L,P). Columbia Co. 58) 1974: 21 Sept (P); 1976: 24 Jan (P) . Dade Co. 6J3) A? Mrs. A.T. Slosson. Desoto Co. 65) 1975: 12 Sept (P,A), 22 Dec (P) . 66) 1975: 22 Dec (L) Dixie Co. 68) 1975: 26 March (L). 69) 1975: 26 March (L) . Flagler Co. 77) 1974: 25 May (P) ; 1975: 5 April (L,P). Glades Co. 95) 1974: 21 March (P) ; 1975: 9 Aug (P) . Hamilton Co. 99) 1975: 23 June (P) , 3 Aug (L,P). Hendry Co. 106 ) 1974: 29 Nov (L,P) ; 1975: 21 March (L,P), 3 July (L,P,A), 31 May (L,P), 9 Aug (P) . Hillsborough Co. 113 ) 1975: 22 March (L,P), 27 May (L,P) , 11 Sept (L,P,A), 21 Dec (L,P) . Lafayette Co. 129 ) 1974: 5 Aug (P) , 12 Oct (P,A); 1975: 17 Jan (L) , 26 March (P) , 28 Dec (P) . 130 ) 1974: 5 Aug (L,P), 12 Oct (P) ; 1975: 26 March (L,P), 23 Aug (L,P), 28 Dec (P) . Lake Co. 170 ) 1974: 11 July (P) , 3 Sept (L) . Levy Co. 135 ) 1974: 3 Aug (L,P), 15 Sept (L,P); 1975: 31 Jan (P) , 23 March (L,P), 1 Sept (P) . 136 ) 1974: 18 June (P). 137 ) 1974: 2 March (P), 27 April (P) , 3 Aug (L,P), 15 Sept (P) , 2 Nov (P) ; 1975: 23 March (P) , 23 Oct (P). 139 ) 1975: 30 Oct (P) , 21 Dec (P,A). 141 ) 1975: 17 May (L.P.A), 8 July (P) , 1 Sept (P) , 30 Oct (P). Madison Co. 151 ) 1974: 8 Aug (P) ; 1976: 14 Feb (P) . Manatee Co. 157 ) 1975: 12 Sept (P) . Polk Co. 182) 1975: 22 March (P) , 21 Dec (L,P).

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-155Putnam Co. 189 ) 1975: 12 Feb (L) . Seminole Co. 202 ) 1974: 21 March (L,P,A), 3 Sept (P) ; 1975: 15 March (P), 4 July (L,P), 31 Oct (P) . 203 ) 1974: 3 Sept (P). Suwanee Co. 208 ) 1974: 17 Sept (P) ; 1975: 24 Oct (P) . Taylor Co. 213) 1974: 14 June (L,P), 5 Aug (L,P), 12 Oct (P) ; 1975: 17 Jan (L,P), 26 March (L,P), 23 Aug (L,P). Union Co. 217 ) 1974: 24 Aug (P) , 9 Oct (P) . Simnlium (Phosterodoros) jonesi Stone and Snoddy Simulium jonesi Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 29 (female, male, pupa, larva). Taxonomy . Stone and Snoddy created a new subgenus, Phosterodoros , in 1969 and S. jonesi was one of eleven new species named at that time. Stone and Snoddy (1969) indicate that the type locality is Fish River, Baldwin Co., Alabama, and that the female holotype with its pupal exuvium and cocoon has been deposited in the U.S. National Museum. Descriptio n. The larva is about 4.5 mm long. The cephalic apotome is yellow with brown spots (Fig. 58). The gular notch is about as wide as long, extends slightly more than halfway to the teeth of the submentum, and may be pointed anteriorly (Fig. 59). The cephalic fans possess 49 to 54 rays each. The anal gills are compound or arborescent and the anal tubercles are well developed and conical. The pupa is 2.5 mm long. The cocoon has convex anterior edges in lateral view and a large aperture on each side anteriorly. The pupal respiratory organ consists of a stout basal filament which tapers distally and nine additional filaments which arise from the basal one (Fig. 60).

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-156ft Figure 58. Cephalic apotome of a S. jonesi larva. Figure 59. Gular notch of a S. jonssi larva.

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-157Figure 60. Respiratory organ of a S. jonesi pupa.

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-158The male is black and velvety in appearance with a pair of triangular, silvery, iridescent spots anterolateral^ on the scutum. The ventral plate in ventral view has the middle portion widened distally and the arms diverge and bear posterior projections. In end view the plate is stout and tapers to a point (Fig. 61). The female has a shiny blackish gray scutum, a shiny black frons, and shiny terminal abdominal tergites. The clypeus is about as wide as long. There are no hairs under the subcosta. The fore legs are darker brown basally on the tibia and femur and more yellowish on the distal ends of the segments. The tarsal claws lack a prominent basal projection. The genital fork is in the shape of a widespread-!' (Fig. 62). Distribution . Stone and Snoddy (1969) mention that early collections of the species now known as S. jonesi were made by CM. Jones in 1954 in Florida and South Carolina and, in addition, they list collection records in Alabama for this species. Life hi story. In Alabama summer eggs are oviposited on grasses and other aquatic vegetation and the egg is believed to be the overwintering stage (Stone and Snoddy, 1969). Stone and Snoddy (1969) indicate egg hatch occurs in late February or March with mature pupae appearing by April and they state at least four generations are completed each year. Habit s. Little is recorded in the .literature on the habits of S. QOiiesi. The lack of a strong basal tooth on the tarsal claw suggests this species is mammalophilic although research in Florida has revealed the females will feed on turkeys in the laboratory.

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-159Figure 61. Terminalia of a male of S. jonesi. Figure 62. Genital fork and terminalia of a female of 5. jonesi.

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-160Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 5.85 m 37.72 cm 4.7 19.6°C (67.3°F) .51 m/sec (1.66 ft/sec) Min: .2 1 3.5 8.3 (47) .15 ( .5) Max: 100 200 + 6.9 28.3 (83) 1.53 (5) Simuliwn jonesi was found in 19 counties at 57 sites (Fig. 63). The double circles used as site designations on the map were reduced to single circles in Alachua county due to site denstiy. From Fig. 63 the restriction of this species to the southern coastal plain and northeastern Florida is apparent. It was not found south of Alachua and Putnam counties. All stages of 3. jonesi were found year round in many streams, indicating seven or more generations per year. This species occurs in reasonable numbers in small flows 1-2 m wide such as Little Hatchet Creek (Site 18), the headwaters of Hatchet Creek (Site 36), and Site 56 and is the predominant species at specific collection sites in large flows like the Blackwater River (163) and the Bay Shoal River (221) which are 20-40 in or more wide. The streams most preferred appear to be moderately sized permanent flows about 5-8 m wide, 15-100 cm deep with trailing vegetation, trapped dead tree leaves, some shade and low pH such as South Prong Black Creek (57) and one of the paratype locations the Fenholloway River (210) (Fig. 64). Alkaline swamp flows such as more typical for S. lakei and its characteristic counties bordering the Gulf are generally avoided by S. jonesi. Simulium jonesi was found in streams of a more alkaline nature such as Deep Creek (191) and Blue Creek (116) but populations were usually light. In most streams S. jonesi immatures were collected from trailing eel grass, other grasses,

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-161rJ *P» Figure 63. Florida collection locations for 5. jonesi.

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-162Figure 64. Site 210, the Fenholloway River, a S. jonesi collection location.

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-163sedges, and other green aquatic vegetation. However in the larger sandy rivers and also the smaller sandy creeks where S. jonesi occurred like Piney Woods Creek (166) larvae and pupae were found most often on trapped dead tree leaves, pine needles, and twigs and limbs pressed by the flow against bridge supports. Hatchet Creek (17), before a major construction project took place in the vicinity, supported heavy populations of S. jonesi in a narrow (1-2 m) , rapidly flowing (.76 m/sec = 2.5 ft/sec), partly shaded section with abundant eel grass and large leafed aquatic vegetation. After this habitat was destroyed the major sampling location was moved upstream about 100 meters into a deciduous and pine woods through which the creek flowed. In this shaded location the stream was a few meters wider, somewhat more shallow and was almost bare of green ve.jitatioti in the flow. However immature populations of S. jonesi well over 1000/. 09 sq m (1 sq ft) of substrate especially during March, April, and August covered twigs, logs, dead tree leaves and pine needles. Sirrrulium jonesi also occurred year round in the torrential Juniper Creek (195) where the current was usually .92-1.2 m/sec (3-4 ft/sec) and S, jonesi was found on small trailing water oak leaves and branches, dead leaves, pine needles and some grass. Sirrrulium jonesi was associated with 15 species of black flies in the streams and rivers of Florida tying S. tuberosum for the most associations. The most frequently associated species was S. tuberosum, which occurred with S. jonesi in sandy, limited vegetation sites and other locations on 253 occasions, and S. slossonae which was regularly collected (107 times) with S. jonesi where stream vegetation was abundant (Table 3). Sirrrulium vereoundum was collected with S. jonesi, on 44 occasions. In more than a dozen sites from the panhandle to northeast Florida

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•164S. jonesi larvae were found to be infected with mennithid nematodes. In the majority of instances infections were noted in less than 10% of the larvae collected at any time of the year. At South Prong Black Creek (Site 157), a good collection site for S. jonesi, a high incidence of mermithids was noted. In five visits infections of small larvae ranged from 11.5% (13 or 113) to 19% (7 of 36) to 26% (6 or 23) and in November 45 of 85 small larvae (53%) were found infected and no medium or large-sized larvae were recovered although a few pupae were located. Larvae infected wit!, tiny white spheres or large white masses in the abdomen, believed to be protozoan parasites, were noted at about a dozen locations but the rates of infection were low (2-4%). In older larvae of S. jonesi the anterior medial head spots were often observed to be weakly developed or absent. This characterisitc was only occasionally noted in other members of the Phosterodoros group where the anterior head spots were normally dark and distinct. Forms believed to be variations of S. jonesi were collected in Rock Creek at Torreya State Park (Site 148) and in the Apalachicola River (145). At the first location Phosterodoros pupae were found with respiratory filaments rising from a thicker basal one similar to the respiratory organ of S. jonesi; however there were only 9 filaments total and filaments 5 and 6 (numbered proximally to distally) branched off a noticeable petiole which in some specimens was at least one-third the length of the filaments themselves. At the Apalachicola River site typical S. jonesi pupae were mixed with other less typical pupae with respiratory organs with 10 filaments. Tnese unusual pupae had a thick and elongate basal filament, but had filaments 5, 6 and 7 rising in a group at the end of a long petiole sometimes equal to the lengths of the filaments themselves.

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-165Petioles which are short or nonexistent in this area of the respiratory organ are more typical of S. jonesi. Simpson et al. (1956) reported finding Simulium species No. 58 (=S. jonesi, Stone and Snoddy, 1969) with other black flies in streams in Florida near two outbreaks of L. smithi in turkeys. During the current research immatures were collected from the Santa Fe River (Site 19) where a large population of S. jonesi occurred in October 1975 and of the reared females one fed on 24 October on a turkey infected with L. smithi in the laboratory and again fed on 27 and 28 October on two clean turkeys but no disease transmissions resulted. Florida collection records for S. jonesi . Alachua Co. 1) 1973: 2 Nov (S,M,L,P), 14 Nov (S.M.L.P), 7 Dec (S,M,P), 13 Dec (S,M,L,P); 1974: 2 Feb (S,M,L,P), 7 March (S,M,L,P), 12 April (S,M,L,P), 5 May (P) , 6 July (S) , 20 July (S,M,P), 29 Aug (S,M,L), 28 Sept (M,L,P), 30 Oct (S,M,L,P), 23 Nov (S.M.P.A); 1975: 10 Jan (S), 12 Feb (S,M,L,P,A), 4 April (S,M,P), 10 May (C) , 21 June (S,M,L,P,A), 15 July (S,M,P), 25 Sept (S) , 9 Nov (S,M,L,P,C); 1976: 21 Jan (S,M,L,P,A). 2) 1973: 11 Oct (P) ; 1974: 30 June (P) , 17 Aug (M,P). 6) 1973: 22 Sept (S,M,L,P,A), 16 Oct (S,M,P), 25 Oct (S,M,P); 1974: 2 Feb (S,M,L,P), 7 March (S,M,P), 2 April (S,P), 18 May (S,M,P), 30 June (S.M.L.P), 30 July (S) , 14 Sept (S,M,L,P,C); 1975: 15 Jan (S), 18 May (S,M), 16 Aug (S) , 8 Oct (S,M,L), 6 Dec (S,M,P,C). 7) 1973: 13 Dec (S.P.A); 1974: 2 Feb (S,M,L), 14 Sept (S,P,C), 19 Oct (P), 16 Nov (S,M,L,P). 17) 1970: 17 Dec (P Anthony); 1971: 7 Jan (P Anthony), 21 Feb (A Anthony) ; 1974: 2 Jan (S,M,L,P), 2 Feb (S,M,L,P,C), 7 March (S,M,L,P), 12 April (S,M,L,P,C,A), 5 May (S,M,L,P,C), 6 June (S,M,L,P,C), 1 July

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-166(S,M,L,P,C), 24 Aug (S,M,L,P,C,A), 28 Sept (S,M,L,P), 4 Dec (C) ; 1975: 11 Jan (S,M,L,P,C), 1 March (S,M,L,P,C), 30 April (S,M,L,P,C), 21 June (S,M,L,P,C), 16 Aug (S,M,L,P,A), 8 Oct (S,M,L,P,A), 6 Dec (S,M,L,P,C). 18) 1974: 12 Jan (L,P), 2 March (P) , 11 April (L) , 30 June (P) , 24 July (P) , 24 Aug (L,P), 22 Sept (L,P), 25 Oct (L,P), 23 Nov (L,P); 1975: 10 Jan (L,P), 21 Jan (P) , 11 May (L,P), 14 June (L), 10 Dec (L,P). 19) 1975: 17 July (S.M.P), 22 Oct (S,M,L,P,C,A) . 19a ) (L,P D.W. Anthony). 20) 1974: 12 Jan (P) , 2 March (M,L,P), 12 April (L,P), 6 July (L,P), 29 Aug (M.L.P.A), 25 Oct (M,P,A); 1975: 15 Jan (M,L,P), 31 Jan (P) , 18 April (L,P), 26 July (P) , 3 Oct (L,P,A). 24) 1974: 9 March (L,P), 24 May (L,P); 1976: 19 Feb (L,P). 36) 1974: 6 July (S,M,P), 24 Aug (C) ; 1975: 15 Jan (S), 4 April (S), 22 May (S) , 22 Oct (S,M,L). 39) 1974: 23 Nov (P) . Bay Co. 40) 1975: 27 March (S,P), 11 June (S,P,C), 6 Sept (S) , 30 Dec (P). Baker Co. 41) 1974: 4 Jan (S,M,L,P), 1 June (S,M,L,P,C), 17 Sept (S,M,P), 6 Nov (S,M,P,C,A); 1975: 1 Feb (S,M,L), 26 April (S.M.L, P,C), 21 June (S,M,L,P,C,A) , 17 Aug (S,M,L,P,C), 14 Nov (S,M,L,P,C). Bradford Co. 42) 1973: 6 Oct (P) . 45) 1974: 21 Sept (L,P), 16 Nov (L), 22 Nov (L,P,A). Calhoun Co. 49) 1975: 27 March (S,M,C), 11 June (S,M,L,P,C), 6 Sept (M,L,P,C), 30 Dec (S) . 50) 1975: 11 June (P) . 52) 1975: 19 April (S,M,P). Clay Co. 56) 1974: 4 Jan (S,M,L,P,C), 23 Feb (S) , 4 May (S,M,L,P,C), 6 June (S.M.P.C), 20 July (S,M,C), 31 Aug (S,M), 25 Oct (S,M,P), 27 Nov (S,M,L,P,C); 1975: 16 Feb (S,M), 8 April (S,M,P), 21 June (S,M,P,C,A), 2 Aug (S,M,L,P), 19 Oct (S,M,L); 1976: 31 Jan (S,M).

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-16757) 1974: 4 Jan (S,M,L,P,C), 23 Feb (S,M,L,P,C), 4 May (S,M,L,P, C,A), 6 June (S,M,L,P,C,A) , 20 July (S,M,P), 21 Aug (S,M,L,P), 25 Oct (S,M,L,P), 27 Nov (S,P,C); 1975: 16 Feb (S,M,L,P,C), 18 April (S,M,L,P,C), 21 June (S,M,L,P,C,A) , 2 Aug (S,M,L,P,C), 19 Oct (S,M,L,P,C,A); 1976: 31 Jan (S,M,C). Duval Co. 70) 1975: 12 March (L,P), 4 May (S,M,P,C), 1 July (S) , 17 Aug (S,M,L,C). Escambia Co. 21) 1974: 18 March (S,M,L), 16 June (S,M,P), 7 Aug (S,M, L,P,C), 14 Oct (S,M,L,P); 1975: 19 Jan (S) , 28 March (S,M,L,P,C), 12 June (S,M,L,P,C), 7 Sept (S.M.P.C), 31 Dec (S,L,C). 74) 1974: 16 June (S,M,L,P), 7 Aug (M) , 14 Oct (S,M); 1975: 19 Jan (S) , 28 March (S,M,P), 12 June (S,M), 7 Sept (S) , 31 Dec (S); 1976: 18 April (S,M,L,P). 75) 1974: 7 Aug (S,M,P), 14 Oct (S,M,L,P); 1975: 28 March (S) , 12 June (S,M,L,P) , 7 Sept (S,M,P,C), 31 Dec (S,M,L,P,C). Gadsden Co. 81) 1974: 17 March (M,L,P). 83) 1974: 17 March (S,M,P); 1975: 18 Jan (M) , 27 March (S,M), 10 June (S,L,P,A). 85) 1975: 30 Dec (S,P,C). 87) 1974: 15 June (S,M,L,P,A), 5 Aug (S,M,L,P,C), 12 Oct (S,M,P); 1975: 18 Jan (S) , 27 March (S,P), 10 June (P) , 5 Sept (M,P,A). 88) 1975: 30 Dec (S) ; 1976: 17 April (P) . 90) 1974: 17 March (S,P), 6 Aug (L,L); 1975: 29 Dec (M) . 91) 1975: 10 June (M), 5 Sept (S) . Holmes Co. 116 ) 1974: 17 March (L,P,C), 15 June (M.L.P), 6 Aug (P,C), 13 Oct (L,P,A); 1975: 19 Jan (L) , 28 March (M,L,P), 11 June (L.P.A), 6 Sept (M,L,P,C,A), 30 Dec (M,L,P); 1976: 18 April (M,L,P,C). Jefferson Co. 124 ) 1974: 12 Oct (S,P), 26 March (S) , 10 June (S) , 23 Aug (S,M), 29 Dec (S) . 125 ) 1974: 5 Aug (S.M.L.P), 12 Oct (S,M,L,P); 1975: 17 Jan (S), 26 March (S,M,L,P), 10 June (S,M,L, P,C,A), 23 Aug (S,M,L,P,C), 29 Dec (S,M,P).

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-168Liberty Co. 142 ) 1974: 15 June (S) , 13 Oct (S,P,A); 1975: 27 March (S,M), 11 June (S,L,C), 24 Aug (S,M,L,P,A). 144 ) 1973: 18 Dec (S,M,L); 1974: 17 March (S,M,L,P,A), 15 June (S,M,L,P), 6 Aug (S,M,L,P,C), 13 Oct (S,M,L,P,A); 1975: 18 Jan (S,M,P,C), 27 March (S,M,L), 11 June (S,P,C), 24 Aug (S.M.L), 30 Dec (S,P,C). 145 ) 1976: 17 April (P,C). 146) 1973: 18 Dec (S) ; 1974: 15 June (S,M,L,P,C), 6 Aug (P,A), 13 Oct (S,M,L,P); 1975: 18 Jan (S,M,L,P,A), 27 March (S,M,L,P,C), 11 June (S) , 24 Aug (S,M,L,P,C,A) , 30 Dec (S,P). 147) 1975: 27 March (S). 148) 1974: 15 June (S), 13 Oct (S,L,P); 1975: 27 March (S,L,P), 11 June (S,M,L), 6 Sept (S,M,L,P). Nassau Co. 159) 1974: 24 Aug (M) ; 1975: 26 Sept (M) . Okaloosa Co. 163) 1974: 18 March (S,M), 16 June (S.P.C), 7 Aug (S,M, L,P), 14 Oct (S,M,L,P,A); 1975: 19 Jan (S,M,L,P,C), 28 March (S,M), 12 June (S.M.L.P), 7 Sept (S.M.P.C), 31 Dec (S,M,P). 164 ) 1975: 29 March (S,M,L,P,C), 6 Sept (S) . 165) 1974: 18 March (S,M,L,P,C), 16 June (S,M), 7 Aug (S,M,C), 14 Oct (S.M.P.A); 1975: 19 Jan (S) , 28 March (S,M), 12 June (S,M,L), 31 Dec (S,M,C). 166) 1974: 18 March (S,M,L,P,C), 16 June (S,M,P), 7 Aug (S,M,L,P), 14 Oct (S,M,P); 1975: 19 Jan (S.M.P), 28 March (S,M,L,P,C), 12 June (S,M,L,P,C), 6 Sept (S,M,L,P,C), 31 Dec (S,M,L,P,C). Putnam Co. 187 ) 1975: 25 Sept (S,P). 189 ) 1974: 17 Aug (P,A). 191 ) 1974: 19 Jan (S.M.P), 14 April (S), 17 Aug (S.M.P.C), 23 Nov (S,M,L); 1975: 5 April (S.M.P.C), 26 June (S,M,L,C), 31 Oct (S,M). Santa Rosa Co. 195 ) 1974: 18 March (M,L,P), 16 June (P) , 7 Aug (L,P, C,A), 14 Oct (L,P,A); 1975: 28 March (L,P), 12 June (M.L.P.A), 7 Sept (M.L.P), 31 Dec (M,P) ; 1976: 18 April (M,L,P). 196 ) 1974: 18 March (S.M.L.C), 16 June (S,M,P), 7 Aug (M,P), 14 Oct (S,M,L,P);

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-1691975: 28 March (S), 12 June (S,L). Taylor Co. 210 ) 1954: 8 April (Larvae, PupaeCM. Jones); 1973: 17 Dec (S,M,L,P); 1974: 16 March (S,M,L,P), 14 June (S,M,L,P), 5 Aug (S), 12 Oct (S,M,C); 1975: 17 Jan (S,M), 26 March (S,M), 10 June (S.M.L.P), 23 Aug (S), 28 Dec (S). 211 ) 1975: 17 Jan (L) . Union Co. 217 ) 1974: 5 Jan (M) , 24 Aug (L,P,A), 9 Oct (L,P); 1975: 1 June (P), 22 July (L) . Walton Co. 220 ) 1974: 18 March (S,M), 16 June (S,M,L,P), 7 Aug (S,M,L,P), 13 Oct (S,M,L,P,A); 1975: 19 Jan (S,P), 28 March (S,P,C), 11 June (S,M,L,P,C), 6 Sept (S,M,P,C), 21 Dec (S,P,C). 221 ) 1974: 18 March (S,M), 16 June (S,M,L,P), 7 Aug (M,C), 13 Oct (S,M,L,P,A); 1975: 19 Jan (S,M,P), 28 March (S,M,L,P,C,A) , 11 June (S,M,L,P,C,A) , 6 Sept (S,M,L,P,C), 31 Dec (L,P,C). 223 ) 1975: 28 March (S,M,L, P,A), 11 June (S,M,L,P), 6 Sept (S,M,L,C). Simulium (Phosterodoros) lakei Snoddy Simulium lakei Snoddy, 1976, J. Georgia Entomol. Soc. 11(2): 173 (larva, pupa, female, male). Taxonomy . Snoddy (1976) described and named S. lakei which is the fourteenth species to be designated in the Phosterodoros group. The holotype male was collected from Blackbird Creek, Blackbird State Forest, Delaware and it with its associated pupal exuvium and cocoon is located in the U.S. National Museum. Description . The older larvae are 5-5.5 mm long. Preserved specimens are brown-gray in color with brown head spots on a yellowbrown head capsule. The pattern of the spots on the cephalic apotcme

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-170is as in Fig. 65. The distal portion of the antenna extends just beyond the cephalic fan stalk. The gular notch is broadly saggittiform (Fig. 66) with faint lateral spots bordering the notch. The cephalic fans contain 57-60 rays. The anal tubercles are conspicuous and the anal gills are arborescent. The pupa is about 3.5 mm long. The anterior lateral openings of the cocoon are usually large. The respiratory filaments number 9 with filaments 1-6 arising in pairs off short petioles and filaments 7-9 (most ventral) rising off a longer base (Fig. 67). The wings of the male are about 2 mm long. The scape and pedicel of the antenna are reddish brown and lighter in color than the black flagellum. The scutum is black and velvety with bright anterior lateral patches (Fig. 68) . The abdomen is also black and velvety with iridescent spots on the side of segments 2, 6, and 7. The middle portion of the ventral plate expands distally and the ventral plate and the distimeres appear as in Fig. 69. The wings of the female are about 2 mm long. The frons is shiny brownish black. The width of the clypeus is about equal to its length. There are no hairs under the subcosta. The scutum is shiny grayish black. The tarsi are light yellow brown. The genital fork appears as in Fig. 70. Distribution . Snoddy (1976), in addition to the holotype location in Delaware, records S. lakei from South Carolina and states this species may extend into western Georgia although it was not found in collections from West Central Georgia. Life history, ecology, habits . Little else is recorded in the literature concerning the biology and ecology of S. takei. Larvae,

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-171Figure 65. Cephalic apotome of a S. lakei larva. Mr Figure 66. Gular notch and hypostomium of a 5. lakei larva.

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-172Figure 67. Pupa and cocoon of S. lakei. Figure 68. Dorsal view of a male of S. lakei.

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-173Figure 69. Terminalia of a male of S. lakei. Figure 70. Terminalia of a female of S. lakei.

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-174pupae and adults (reared) were obtained in Delaware from early May to late October and larvae and pupae were found in South Carolina from late March through August (Snoddy, 1976) . Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 9.93 m 54.61 cm 6.01 20.8°C (69.4°F) .49 m/sec (1.62 ft/sec) Min: .2 1.7 3.75 7.2 (45) .15 ( .5) Max: 100 1000 7.55 29.4 (85) 2 (6.66) Based primarily on the similarity between the structures of Florida specimens and the description published in Snoddy (1976) for the pupal respiratory organ and confirmation of determinations by Dr. E.L. Snoddy of submitted Florida specimens this species has been called S. lakei, with the observed morphological variations listed below. Specimens have been collected from 45 locations in 24 counties (Fig. 71). This species has not been positively identified from collections in Florida west of Jefferson County but occurs widely in northeastern Florida and ranges south to west of Lake Okeechobee. This species is mulivoltine and in permanent streams larvae and pupae may be found all year long. S. lakei occurs in small temporary flows such as Otter Creek (Site 135, Fig. 72), Rutland Creek (Site 206) and Rocky Creek (213); smallish permanent flows such as Joshua Creek (65), a flow to Newnan's Lake (18), Site 141, and Howell Creek (202); and in larger permanent flows such as the Santa Fe River at Oleno State Park (58) , Blackwater Creek in Hillsborough Co. (113), the Waccasassa River (136, 137) and the Withlacoochee River (151). At least some of these flows are spring fed being characteristically near neutrality or just below in pH reaction and all are

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-175Figure 71. Collection locations for 5. lakei in Florida.

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-176Figure 72. Site 135, Otter Creek, a collection site for S. lakei.

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-177moderately to very swift flowing with currents at constrictions formed by boulders in the Santa Fe River and flows pouring off the concrete at Howell Creek reaching 1.2-1.8 m/sec (4-6 ft/sec) or more. Other collection sites such as Fisheating Creek (95), Blackwater Creek in Lake Co. (170) and Rice Creek (189) were slower flowing and yielded smaller populations of S. lakei. This species was also found to occur year round in Little Haw Creek (77) which was fairly deep and flowed about .46 m/sec (1.5 ft/sec) and Lochloosa Creek (21) which was fairly shallow and slow flowing (.3-. 46 m/sec = 1-1.5 ft/sec); both of these streams were more acidic than is typical for the habitats of this species and averaged around 4.5 pH reaction. The substrate for most sites was sand with some pebbles, rocks, or boulders occurring in the flow. Immatures were typically found attached to a wide range of green aquatic, trailing and trapped vegetation including grasses, sedges, leaves of cattail, smartweed, pickerelweed, Hemlock andwater willow, twigs, dead tree leaves, pine needles and algae or moss on rocks. On eel grass pupae were found often situated on the middle of the blade parallel to the long axis and facing the blade tip. On willow and other tree leaves pupae frequently attached near the outer leaf edge near the serrations parallel with the main rib and were found on both sides of the leaves. The largest populations of immatures were encountered usually during the cooler part of the year from November through April; however, heavy populations were noticed in some permanent flows at other times such as during August in Kettle Creek (Site 130). Simulium lakei was found in Florida streams and rivers at least once with nine other black fly species (Table 3 )• Most frequently S. lakei was associated with the other Phosterodoros species, S. taxodium

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-178(150 associations) and S. jenningsi (92 associations). Sirrrulium slossonae and S. tuberosum were next most commonly found with S. lakei. In September 1975 a reared 5. lakei fed on a turkey in the laboratory. A number of S. Phosterodoros species females which are difficult to separate without associated pupae and may have included females of S. lakei were captured in Manitoba traps at Fisheating Creek and elsewhere and fed on turkeys in the laboratory. However since this species lacks the tooth on the tarsal claw typica] of ornithophilic species, it probably prefers mammals such as the scrub cattle which ranged near many of the breeding sites like Lochloosa Creek (22). Morphological variations have been observed in Florida specimens of S. lakei compared to specimens illustrated from other areas in Snoddy (1976). Some of the Florida larvae display a gular notch which is wider and less elongate than pictured. There appear to be about 76 rows of anal hooks on the larva rather than 56-58 rows. The dorsal most respiratory filament on a number of S. lakei pupae, locally, rises vertically and abruptly proceeds anteriorly at a much sharper angle than the gentle curve drawn. The most ventral filament rather than separating from the common stem of filaments 7 and 8 far out from the base rises very close to the base of the petiole of 7 and 8 and the main organ stem. Many pupae have been collected with 8 filaments on one side and 9 on the other or with a 9 and 10 filament variation. The feaale and male terminalia generally agree with figures for S. lakei. Florida collection records for S. lakei Alachua Co. 1) 1974: 12 April-(L,P). 7) 1974: 2 Feb (P). 17) 1975: 16 Aug (L). 18) 1974: 12 Jan (L,P), 2 March (L.P), 11 April (L,P) 25 May (L,P), 30 June (P) , 24 July (l.,F), 24 Aug (L,P) , 22 Sept

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-179(L,P), 25 Oct (L,P), 23 Nov (L,P); 1975: 10 Jan (L,P), 31 Jan (P) , 8 April (L,P), 11 May (L,P,A), 14 June (P), 17 July (P) , 1 Sept (L,P,A), 17 Oct (L,P,A), 10 Dec (L,P,A). 20) 1974: 12 Jan (P) , 2 March (L,P), 6 July (P), 29 Aug (L,P,A), 23 Nov (P) ; 1975: 15 Jan (L,P), 31 Jan (L,P), 18 April (L,P), 26 July (L,P), 3 Oct (L,P). 21) 1974: 2 March (L,P), 11 April (L,P), 30 July (P) , 14 Sept (L,P), 30 Oct (P); 1975: 24 Jan (L,P), 4 April (L,P), 27 Sept (L,P) ; 1976: 21 Jan (P), 29 Jan (P) . _22) 1974: 15 Aug (L,P), 28 Aug (L,P), 12 Sept (P), 7 Dec (L,P) ; 1975: 24 Jan (L.P.A), 4 April (L,P), 29 April (L,P), 5 Aug (L,P), 30 Aug (L,P), 17 Oct (L,P,A); 1976: 6 March (L,P). 24) 1974: 9 March (P) , 25 May (L,P); 1975: 18 May (P) ; 1976: 19 Feb (L,P). 39) 1974: 25 Oct (P) , 23 Nov (P) ; 1975: 26 July (P) . Bradford Co. 45) 1974: 4 May (P) , 21 Sept (P) , 16 Nov (L) ; 1975: 22 Nov (P,A). Columbia Co. 58) 1974: 23 Feb (P) , 27 April (P) , 21 Sept (L,P), 6 Nov (L,P); 1975: 12 Jan (L,P), 26 April (L,P), 21 June (P) , 17 Aug (L,P), 22 Oct (L). 59) 1974: 27 April (P) , 6 Nov (P) . Desoto Co. 65) 1974: 29 Nov (P) ; 1975: 22 March (L,P,A), 12 Sept (L,P), 22 Dec (L,P). 66) 1975: 22 Dec (P) . Dixie Co. _68) 1975: 26 March (P) , 25 Aug (S,P). 69) 1975: 26 March (P). Flagler Co. 77) 1974: 26 Jan (L,P), 25 May (L,P), 31 Aug (L,P), 7 Dec (L,P); 1975: 5 April (L,P), 9 July (L,P), 31 Oct (P) . Glades Co. 95) 1975: 28 July (P) , 9 Aug (L,P). Hamilton Co. 99) 1974: 8 Aug (L,P) ; 1975: 1 Feb (L) , 26 April (L,P), 23 June (P) , 3 Aug (L,P) , 24 Oct (P).

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-180Hardee Co. 104 ) 1975: 21 Dec (P) . Hendry Co. 106 ) 1974: 29 Nov (L,P,C); 1975: 31 May (L,P), 3 July (M,L,P), 9 Aug (P), 16 Oct (L). Hillsborough Co. 113 ) 1975: 22 March (L,P), 27 May (L,P), 11 Sept (M,L), 21 Dec (L,P,A). Jefferson Co. 125) 1974: 5 Aug (L) . Lafayette Co. 129 ) 1974: 16 March (L,P) , 14 June (L,P,C), 5 Aug (L) , 12 Oct (L,P,C); 1975: 17 Jan (L) , 26 March (L,P), 23 Aug (L,P), 28 Dec (P). 130 ) 1974: 14 June (L,P), 5 Aug (L,P), 12 Oct (L,P); 1975: 17 Jan (L,P), 26 March (L,P), 10 June (L,P), 23 Aug (L,P), 28 Dec (P). Lake Co. 170 ) 1974: 28 Nov (P) . Levy Co. 135 ) 1973: 7 Oct (F) ; 1974: 2 March (L,P), 27 April (L,P,C), 3 Aug (L,P), 15 Sept (L,P) ; 1975: 21 Jan (L,P), 23 March (L,P), 17 May (P) , 1 Sept (L,P,A), 30 Oct (L,P,C), 21 Dec (L,P) . 136 ) 1974: 2 March (L,P), 18 June (P) , 3 Aug (L) , 15 Sept (L,P,C), 2 Nov (L,P,A); 1975: 31 Jan (L,P), 23 March (L,P,A), 17 May (P) , 30 Oct (L,P). 137 ) 1973: 29 Dec (L,P); 1974: 2 March (L,P), 27 April (L,P), 3 Aug (L,P), 15 Sept (L,P), 2 Nov (L,P,A); 1975: 31 Jan (L,P), 23 March (L,P,A), 5 July (L,P,C), 1 Sept (L,P), 30 Oct (L, P,A). 138 ) 1975: 31 Jan (L,P). 139 ) 1975: 23 March (P), 30 Oct (L,P), 21 Dec (L,P). 141) 1975: 23 March (L,P), 17 May (L,P), 8 July (L,P), 1 Sept (L,P,A), 30 Oct (L,P). Madison Co. 151 ) 1974: 10 Nov (L,P); 1975: 1 Feb (P) , 26 April (L,P), 23 June (P) , 3 Aug (P) ; 1976: 14 Feb (P) . Manatee Co. 157 ) 1975: 12 Sept (L,P), 22 Dec (L,P). Polk Co. 181) 1974: 29 Nov (L,P,A); 1975: 22 March (L,P,C), 21 Dec

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-181(L,P,A). 182 ) 1975: 11 Sept (P), 21 Dec (L,P). Putnam Co. 186 ) 1974: 25 May (L) , 23 Nov (P) . 189 ) 1974: 19 Jan (P), 14 April (P,C), 17 Aug (L,P,C), 6 Oct (P) , 23 Nov (P) ; 1975: 12 Feb (L), 31 Oct (P) . Seminole Co. 202 ) 1974: 21 March (L,P), 12 May (P) , 11 July (L,P), 3 Sept (L,P), 28 Nov (L,P,C); 1975: 15 March (L,P,C), 27 April (L,P,A), 4 July (L,P), 31 Oct (L,P). 203 ) 1974: 3 Sept (P) , 28 Nov (L,P,C); 1975: 15 March (L,P,C), 31 Oct (L) . Sumter Co. 206 ) 1973: 21 Nov (M,P) . Suwanee Co. 207 ) 1973: 6 Nov (P) . 208 ) 1974: 8 Aug (L) , 17 Sept (P,A), 10 Nov (P); 1975: 1 Feb (L,P,A), 26 April (L,P,A), 23 June (P), 24 Oct (L,P); 1976: 24 Jan (L) . Taylor Co. 213 ) 1974: 14 June (P) , 5 Aug (P), 12 Oct (L,P,A); 1975: 26 March (L,P), 23 Aug (L,P). Union Co. 217 ) 1974: 23 Feb (L,P), 4 May (P) , 6 July (L,P); 1975: 22 July (L,P,A). Simulium (Phosterodoros) notiale Stone and Snoddy Simuliwn notiale Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 40 (female, male, pupa). Taxonomy . A male was described and named as the holotype for this species. The type locality is Meadows Mill, Lee Co., Alabama. The holotype male with associated pupal exuvium and cocoon has been deposited in the U.S. National Museum (Stone and Snoddy, 1969). Description . Stone and Snoddy (1969) did not positively identify the larvae. During the current research S. notiale larvae were observed

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-182to have the following characteristics: the length of the larvae was 5-5.75 nun; the head capsule appeared yellow brown with dark brown head spots and the abdomen was greenish-brown which faded to gray in alcohol; the posterior lateral head spots were not slanted but positioned horizontally on the epicranial plate (Fig. 73); the head capsule appeared fairly wide with convex margins when viewed dorsally; the gular notch was broad, extended over half the distance to the submental teeth, was bordered by a pair of dark oval spots along its length and was either broadly rounded or more pointed anteriorly (Fig. 74); the anal tubercles were inconspicuous. The pupa is 2.5-3 mm long. The respiratory organs each consist of 6 filaments in 3 pairs on short petioles with the filaments' length being about one-third to one-half the length of the pupa. The cocoon is joined anterioventrally (Fig. 75). The male is velvety black with a pair of large triangular iridescent areas laterally on the forward part of the scutum. The velvety black area between the iridescent spots is broadly expanded anteriorly and expands posteriorly and meets a broad more dull, gray region at the rear of the scutum (Fig. 76). The abdomen is velvety black with iridescent patches along the side. Terminalia are as in Fig. 77. The female bears wings about 2.5 mm long. The hairs are dark on the stem vein and costa. The frons is shiny black. The clypeus is thinly gray pollinose and slightly longer than wide. The scutum is shiny dark brown with a pair of pale gray spots anteriorly. The fore tibia bear elongate bright white patches that extend threefourths of the length of the tibia. The anterior abdominal tergites are velvety black while the posterior four tergites are shiny black. Terminalia are as in Fig. 78.

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-183Figure 73. Head spots of a S. notiale larva. Figure 74. Gular notch of a S. notiale larva.

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-184Figure 75. Pupal exuvium and cocoon of S. notiale. Figure 76. Scutum of a male of 5. notiale.

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-185/ i Figure 77. Terminalia of a male of 5. notiale. Figure 78. Terminalia of a female of S. notiale.

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-186Distribution . Simuliwn notiale has been reported from Alabama, South Carolina and Virginia (Stone and Snoddy, 1969). Life history . Stone and Snoddy (1969) found S. notiale only in the early spring and suggest that the eggs overwinter and that the species is univoltine. Ecology . In Alabama pupae of S. notiale were collected in mediumsized streams with high quality water exhibiting a temperature of 1316°C and a pH of 7.1-7.2. Pupae were found in waterfalls at the top of a dam attached to sticks in a flow of .76-. 92 m/sec (2.5-3 ft/sec) where S. venustum also occurred. Habits . The habits of this black fly are unknown. Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 35 m; 5.5 m 4.3 cm; 18.74 cm 4.51 18.3°C (65°F) 1.16 m/sec (3.79 ft/sec) Min: 34 ; 4 3 ; 5 4.45 9.4 (49) .67 (2.2) Max: 36 ; 6.5 5 ; 25 4.65 25.6 (78) 1.53 (5) Simulium notiale was found in one county, Gadsden, at two sites adjacent to each other, Sites 88 and 89, in Chattahoochee, Florida, during April, August and December (Fig. 79). All stages were present during each month; however, the smallest populations were found during December and the largest during August. In Florida this species appears multivoltine and capable of completing generations most all of the year. In the stream parameter table the width and depth figures are listed for each site separately. At Site 88 (Fig. 80) S. notiale immatures were collected from dead leaves, pine needles, and small amounts of grass at the crest of a rounded, concrete, power station dam about 35 m long and 6.1 m (20 ft)

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-187r/ J*** Figure79. Collection locations for S. notiale in Florida,

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-188Figure 80. Site 88 at Chattahoochee where S. notiale immatures were found.

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-189high. The water poured at high velocity a few centimeters deep from a large impounded lake. No S. notiale immatures were found attached to the concrete which was coated with a thin layer of green algae. At Site 89, Mosquito Creek, immatures were recovered a short way downstream from a second, smaller dam in a rapidly flowing, moderately deep stream about 5 m wide which flowed over exposed yellow, slippery limestone. Little vegetation occurred in the flow but black snails were abundant. Hardwoods and pines crowded in on the stream. In August large populations of S. notiale were found attached to outcrops of the yellow rocklike substrate, moss, pine needles, green trailing tree and bush leaves, dead leaves, and a few blades of grass. The pH of both sites was below 5.0 and the immatures were found in water from 9.4°C to 25.6°C (49-78°F) . Simulium notiale was collected with S. tuberosum, S. jonesi, S. decorum and S. vereoundum (Table 3 ) . Two reared females each fed once on a turkey in the laboratory during August but died within two days of the feedings. Florida collection records for S. notiale . Gadsden Co. 88) 1975: 24 Aug (S,M,L,P,C), 30 Dec (S,M); 1976: 17 April (M,P). 89) 1975: 8 Aug (S,M,L,P K. Tennessen), 24 Aug (S,M,L,P,A), 30 Dec (S,M,P,C,A); 1976: 17 April (S,M,L,P,A). Sirmxlium (Phosterodoros) nyssa Stone and Snoddy Simulium nyssa Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 42 (female, male, pupa). Taxonomy . A male was described and designated the holotype of S. nyssa by Stone and Snoddy (1969). The type locality is Meadows Mill,

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-190Lee Co., Alabama. The holotype male has been deposited in the U.S. National Museum. Description . Stone and Snoddy (1969) state that the larva is apparently not separable from S. jonesi or 5. dixiense without examination of the respiratory histoblast. No larvae recognizable as S. nyssa were collected in the current research. The pupa is 3 mm long in a typical Phosterodoros cocoon with lateral apertures. The pupal respiratory organ consists of 10 thin filaments. The dorsal or posterior 4 filaments are shorter than the other filaments and arise from short petioles or are nearly sessile. The remaining 6 filaments consist of two pairs each with a long petiole and a third filament which rises off the petiole of each pair (Fig. 81). The male has a gray pollinose clypeus, brown palpi, and a black scutum with a pair of iridescent spots near the anterior margin. The dark area between the spots widens anteriorly and posteriorly. The abdomen is deep reddish brown with the usual silvery pollinose areas (Stone and Snoddy, 1969). The female has wings which are 1.75-2.3 mm long with dark stem vein hairs. The frons is shining black; the clypeus is lightly gray pollinose, slightly longer than wide. The scutum is shiny black with a pair of pale gray spots anteriorly and thin recumbent coppery hair. The abdomen is dark with a yellow to coppery basal fringe and shiny terminal terga (Stone and Snoddy, 1969). Distribution . Stone and Snoddy (1969) list records for S. nyssa from Maine and Connecticut in the north, Oklahoma, Arkansas, and Texas out west and Virginia, Mississippi, Missouri, Kentucky, Louisiana, Alabama, South Carolina, and Florida in the south.

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-191Figure 81. S. nyssa pupa and cocoon.

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-192Life history . In Alabama four or five generations are completed each year with overwintering occurring in the egg stage (Stone and Snoddy, 1969). Sleeper (1975) reports three or four generations for S. nyssa in Maine. Ecology . Stone and Snoddy (1969) report S. nyssa is commonly collected on river weed, Podostemon ceratophyllum, in shallow rapids (.761.06 m/sec = 2.5-3.5 ft/sec) in large streams in association with S. underhilli and in small streams with S. snowi. Immatures prefer water temperatures 13-28°C and have been collected in water with a pH reaction of 6.8-7.3. Sleeper (1975) reports that swift, cool, pure mountain streams sustain S. nyssa in Maine. Habits . Stone and Snoddy (1969) observed females believed to be S. nyssa ovipositing on the face of a dam by flying up and down and tapping their abdomens to the trickle that slowly flowed over the dam surface. Females are reported to be primarily annoying to man which they seldom bite but are said to be serious pests of cattle, attacking the ears, stomach, genitalia and other regions. Sleeper (1975) reports S. nyssa is a vicious man-biter in Maine. Florida observations . Stream Parameters Width Depth pH Temperature Velocity 6 m 30-90 cm 6.75 14.2°C (57.5°F) .85 m/sec (2.8 ft/sec) Simuli-wn nyssa is reported from one site in each of two counties in Florida (Fig. 82). The record for Orange County refers to pupae collected in January 1947 from the Big Econlockhatcb.ee River in Orlando. Three pupae were examined at the U.S. National Museum that were labeled

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-193Figure 82. Collection locations for 5. nyssa in Florida.

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-194Figure 83. Site 116, Blue Creek, where S. nyssa was collected.

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-195S. nyssa but were observed to have only 9 filaments in the respiratory organ similar to S. lakei which Snoddy (1976) recently described. If this is the Florida record referred to in Stone and Snoddy (1969) other specimens not examined in the current study must be on hand or the record is questionable. In this research only one pupa which appears to be S. nyssa was collected, and that at Blue Creek, Site 116 (Fig. 83). The collection was made on 19 January and the pupa was attached to a thin twig. The stream parameters for Blue Creek at the time of the collection are presented above. The stream substrate consisted of planks, logs, twigs, debris and sand. Little green vegetation except trailing tree leaves occurred in the section of the stream sampled. The pupa was collected with specimens of S. jonesi and S. tuberoswn. The Florida S. nyssa pupa has the solitary filament of each lower or anterior triplet on the respiratory organ rising ventrally off the long petioles of the other .filaments instead of dorsally as figured for S. nyssa in Stone and Snoddy (1969). However specimens from Oklahoma observed at the National Museum and labeled S. nyssa appear like the Florida specimen. Florida collection records for S. n yssa. Holmes Co. 116) 1975: 19 Jan (P) . Orange Co. 171 ) 1947: 9 Jan (P H.K. Gouck). Simu Hum (Pkoeterod or-os) taxodium Snodd y an^JBeshear Simulium (Simulium) taxodium Snoddy and Beshear, 1968, J. Georgia Entomol. Soc. 3(3): 123 (female, male, pupa, larva). Taxonomy. Snoddy and Beshear (196S) designated a female with its i

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-196associated pupal exuvium and cocoon as the holotype. The type locality is Chickasawhatchee Creek at Highways 37 and 216 in Baker County, Georgia. The holotype is deposited in the U.S. National Museum. Paratypes are located in the museums of the University of Georgia at Athens and Experiment, Georgia. Description . Fully developed larvae are 4.5 mm long. The head capsule is yellow brown with brown head spots. The abdomen of preserved specimens is yellow brown to gray brown in color. Snoddy and Beshear (1968) state the gular notch is bulbous and broadly rounded anteriorly, that each caphalic fan contains 38-42 rays and that there are 76-82 rows of anal hooks (Fig. 84 and 85). The pupa is 2.5-3 mm long. The respiratory organs each consist of 8 filaments arranged in four pairs. The petiole of the third pair, from the dorsal, is the longest (Fig. 86). The cocoon is tightly woven, slipper-shaped and bears a pair of large anterior-lateral openings. The wing of the male is about 2 mm long. Large bright silvery patches occur on the forward-lateral areas of the scutum and the posterior is silvery with the remainder of the scutum being velvety black. The stem vein hairs are dark brown. The abdomen is velvety black with silver patches along the side. The distimeres are elongate, taper distally and end with one large terminal spine. The male terminalia in ventral view appear as in Fig. 87. The female wings are 2-2.1 mm long. The frons is shiny dark brown as are the terminal abdominal tergites. The clypeus is gray pollinose. The scutum is blackish gray and shiny with thin gray pollinosity laterally and sparce, small golden hairs. The subcosta is bare. The cerci are broadly rounded distally, bear many setae and in lateral view appear

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-197Figure 84. Cephalic apotome and head spots of a S. taxodium larva. Figure 85. Gular notch of a S. taxodiiun larva.

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-198Figure 86. Pupal exuvium and cocoon of S. taxodium. % Figure 87. Male terminalia of S. taxodium.

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-199> Figure 88. Female terminalia of S. taxodium.

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-200as one half of an oval almost as wide as the anal lobe. The genital fork is as in Fig. 88. Distribution . Simulium taxodium is reported from southwestern Georgia (Snoddy and Beshear, 1968). Life history . Little has been published on the life history of this species. Mature larvae and pupae were found to occur during early March but were absent in July and August (Snoddy and Beshear, 1968). Ecology . Snoddy and Beshear (1968) report that bald cypress, Taxodium dietiahum Rich., is common where this black fly breeds. Habits . Nothing has been seen in the literature concerning the habits of this species. Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 11.65 m 58.2 cm 6.09 20.4°C (68.7°F) .54 m/sec (1.77 ft/sec) Min: .15 2 3.75 7.2 (45) .15 ( .5) Max: 100 1000 7.55 29.2 (84.5) 1.78 (5.85) Simulium taxodium has been found in 23 counties from west to south Florida at 39 locations (Fig. 89). This species has multiple generations each year and larvae and pupae have been collected during each month of the year. Simuliwn taxodium has been collected from a range of streams including streams with low pH reactions (4.3-4.5) such as Hatchet Creek at Site 20 and Site 99, the Alapaha River a large temporary flow which is usually just a dry white sand river bed in November or December of each year. Most preferred are streams with a pH reaction close to neutral (6.4-7.2) such as the intermittent, small Otter Creek (Site 135) which has been discovered not flowing during June and July as well as

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-201Figure 89. Collection locations for S. taxodium in Florida.

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-202October and November and the permanent, larger, spring-fed Ichetucknee River (Site 59), both with abundant aquatic vegetation. Other flows with fairly neutral pH but little trailing vegetation, such as the Santa Fe River at Oleno State Park (Site 58) with moss-covered boulders a prominent feature in the flow and Blue Creek (Site 116) with considerable trapped twigs, leaves and other debris, consistently yielded S. taxodium. Cypress stands were notably present at the Ichetucknee River and Santa Fe River collection locations. Populations were found, generally, to be the largest when and where the flow was most swift, usually in the range .61-1.22 m/sec (2-4 ft/sec), which varied throughout the year in the various streams. At the Ichetucknee River (Fig. 90) S. taxodium was the predominant and almost exclusive black fly species present and was found year round on eel grass. At other locations S. taxodium immatures were collected from grass trailing from the banks, a wide variety of aquatic vegetation, vine, holly and water willow leaves, dead tree leaves, pine needles, twigs and rocks. Simulium taxodium was associated with nine other black fly species in the streams and rivers (Table 3 ) . Most frequently collected in association were S. takei and S. jenningsi, two other members of the subgenus Phosterodoros . The widespread species S. slossonae and S. tuberosum were next most frequently collected with S. taxodium, followed by another Phosterodoros species, S. jonesi. Florida specimens of S. taxodium differed from the descriptions in Snoddy and Beshear (1968) in a few ways. The larvae frequently had broad but pointed gular notches and the number of rays in each cephalic fan numbered between 50-60 rather than 38-42. The third petiole of the pupal respiratory organs was the longest of the four petioles but was

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-203Figure 90. The Ichetucknee River, a S. taxodiwn collection site.

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-204usually only 1/5 or 1/4 the length of the filaments themselves rather than the described length of 1/2 that of the filaments. The ventral plate was observed to expand distally in ventral view rather than remain fairly parallel-sided to the end and the plate in end view was not visible as a narrow sharp pointed structure but more rounded and tipped with a small knob. One reared female of S. taxodium appeared to feed on a turkey in the laboratory as evidenced by a tiny feeding mark on the skin. When dissected three days later no blood was noticed in the gut of the fly and the ovaries lacked developed eggs. Florida collection records for S. taxodium. Alachua Co. 18) 1974: 12 Jan (L,P,A), 2 March (P) , 11 April (L,P) , 30 June (P), 24 July (L,P), 24 Aug (L,P) , 22 Sept (L,P), 25 Oct (L,P), 23 Nov (L,P,A); 1975: 10 Jan (P) , 31 Jan (L,P), 8 April (P,A), 11 May (P), 14 June (L) , 1 Sept (P) , 17 Oct (P) , 10 Dec (L,P). 20) 1974: 12 Jan (L,P), 2 March (L,P), 12 April (L,P), 6 July (P), 29 Aug (P) ; 1975: 15 Jan (L) , 31 Jan (L,P), 18 April (L,P), 26 July (P), 3 Oct (P) . _21) 1974: 11 April (L,P), 14 Sept (L,P), 30 Oct (P); 1975: 24 Jan (L,P), 4 April (L,P), 27 Sept (L,P); 1976: 21 Jan (L,P), 29 Jan (L,P). TT) 1974: 28 Aug (L,P), 12 Sept (L), 7 Dec (L,P,A); 1975: 24 Jan (P) , 4 April (L,P) , 29 April (P), 5 Aug (P,A), 30 Aug (P) , 17 Oct (P,A); 1976: 6 March (P). 24) 1974: 9 March (P) , 25 May (L,P), 18 May (P) . 39) 1975: 26 July (L,P). Bradford Cc. 45) 1974: 4 May (P) , 16 Nov (P,A); 1975: 22 Nov (P) . Calhoun Co. 50) 1975: 11 June (P,A).

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-205Columbia Co. 58) 1973: 25 Aug (L,P); 1974: 5 Jan (L,P,A), 23 Feb (L,P,C), 27 April (L,P,A), 1 July (L,P), 3 Aug (P) , 21 Sept (P), 6 Nov (L,P); 1975: 12 Jan (L,P), 26 April (L,P,A), 21 June (L,P), 17 Aug (L,P), 22 Oct (P,C,A); 1976: 24 Jan (L,P). 59) 1973: 23 Sept (S.M.L.P.A), 6 Nov (S,M,L,P), 19 Dec (S,M,L,P); 1974: 23 Feb (S,M,L,P), 27 April (S,M,L,P), 7 May (S,M,L,P,A), 29 June (S,M,L,P), 3 Aug (S,M,L,P), 12 Aug (S,M,L,P,A), 21 Sept (S,M,L,P), 6 Nov (S,M, L,P); 1975: 12 Jan (S,M,L,P), 29 March (S,M,L,P), 21 June (S,M), 22 July (S,P,C), 27 Sept (S,M,L,P,C,A) , 22 Nov (S,M,L,P). 62) 1975: 1 Nov (S,P). Desoto Co. 65) 1974: 29 Nov (P,A); 1975: 22 March (P,A), 12 Sept (L) , 22 Dec (P). 66) 1975: 12 Sept (P) , 22 Dec (P) . Dixie Co. 69) 1975: 26 March (L) . Duval Co. 70) 1975: 12 March (L,P). Flagler Co. 77) 1974: 7 Dec (P) ; 1975: 5 April (P). Glades Co. 95) 1975: 17 June (P,C). Hamilton Co. 99) 1974: 8 Aug (L) ; 1975: 26 April (L) , 23 June (P) , 3 Aug (L,P), 24 Oct (P,A). Hardee Co. 104 ) 1975: 21 Dec (L) . Hillsborough Co. 113 ) 1975: 22 March (L,P), 27 May (L,P,A), 11 Sept (L,P), 21 Dec (L,P). Holmes Co. 116 ) 1974: 17 March (M,L), 6 Aug (P) , 13 Oct (L,P); 1975: 28 March (L,P) , 11 June (P) , 6 Sept (M) , 30 Dec (P) . Lafayette Co. 129 ) 1974: 16 March (P,A); 1975: 26 March (L,P), 23 Aug (P), 28 Dec (P,A). 130 ) 1974: 12 Oct (L,P); 1975: 26 March (P) , 10 June (P) , 23 Aug (P) , 28 Dec (P) . Lake Co. 170 ) 1975: 3 July (P) .

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-206Levy Co. 135 ) 1973: 7 Oct (P) ; 1974: 2 March (L) , 27 April (L,P,C), 3 Aug (L,P), 15 Sept (P) ; 1975: 31 Jan (P,A), 23 March (L.P.A), 17 May (P) , 1 Sept (L,P), 30 Oct (P) , 21 Dec (L,P). 136 ) 1973: 30 Oct (P), 29 Dec (L) ; 1974: 2 March (L,P), 18 June (P) , 15 Sept (P), 2 Nov (L,P); 1975: 31 Jan (P) , 23 March (L) , 1 Sept (P) , 30 Oct (P). 137 ) 1973: 29 Dec (L,P); 1974: 2 March (L,P), 27 April (L,P), 3 Aug (L,P), 15 Sept (L,P), 2 Nov (P) ; 1975: 31 Jan (P) , 23 March (L,P), 1 Sept (L,P), 30 Oct (P). 138) 1975: 31 Jan (P) . 139 ) 1975: 11 Sept (L) , 30 Oct (L,P), 21 Dec (L,P). 141 ) 1975: 17 May (L,P), 8 July (P) , 1 Sept (P) , 30 Oct (L,P) . Madison Co. 151 ) 1973: 19 Dec (P) ; 1974: 10 Nov (L,P); 1975: 1 Feb (L), 25 April (L,P), 23 June (L,P,A), 3 Aug (P,A), 24 Oct (L) ; 1976: 14 Feb (P) . Manatee Co. 157 ) 1975: 12 Sept (L,P), 22 Dec (L,P). Polk Co. 181 ) 1974: 29 Nov (L) ; 1975: 22 March (P) , 21 Dec (P,A). 182 ) 1975: 11 Sept (P) , 21 Dec (L,P). Seminole Co. 202 ) 1974: 12 May (L) ; 1975: 15 March (P) . Suwanee Co. 207 ) 1973: 23 Sept (P,A). 208 ) 1974: 10 Nov (L) ; 1975: 1 Feb (L), 26 April (L,P); 1976: 24 Jan (P) . Taylor Co. 213 ) 1974: 14 June (P) , 5 Oct (P) ; 1975: 17 Jan (P) , 26 March (L,P). Union Co. 217 ) 1974: 4 May (L,P), 6 July (L,P), 24 Aug (P) , 9 Oct (P) , 6 Nov (P); 1975: 22 July (P,A). Simuliim (Psilozia) vittatum Zetterstedt Simulia vitvata Zetterstedt, 1838, Insecta Lapponica, 1838-1840: 803 (female) .

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-207Simuliwn tribulation Lugger, 1897, Minn. State Entomol. Rep. 2: 179 (female, male, larva, pupa). Sirrrulium glauaum Coquillett, 1902, Proc. U.S. Nat. Mus. 25: 97 (male). Simulium venustoides Hart, 1912, in Forbes. 111. State Entomol. Rep. 27: 42 (male only). Psilozia groenlandica Enderlein, 1936, Gesell. naturf. Freunde, Sitzber. 114 (female). Simulium asakakae Smart, 1944, Roy. Entomol. Soc. London, Proc. (B) 13: 131. Simulium vittatum — Stone, 1964, Conn. State Geol. and Natur. Hist. Surv. Bull. 97: 40 (female, male, larva, pupa). Simulium vittatum— Stone and Snoddy, 1969, Auburn Univ. Agr. Exp. Sta. Bull. 390: 29 (female, male, larva, pupa). Taxonomy . Zetterstedt (1838) first described a female of this species from Greenland. Stone (1964) indicates the holotype is in the University of Lund, Sweden. Peterson (1965) described the female specimen at the University of Lund and designated it as the lectotype in the event other of Zetterstedt ' s specimens materialize. Lugger (1897) described a male, larva and pupa for this species under the name S. tribulatum. Davies et al. (1962) suggest that this widely distributed black fly may be a complex of two species. Pasternak (1964) examined the larval salivary gland chromosomes of S. vittatum specimens from Alaska, Canada, and New England and concluded that no sibling species existed but that the species was very polymorphic due to exploitation of many niches. Description . Wu (1930) describes S. vittatum eggs as .25 mm long, .15 mm broad and .14 mm high. The larvae are 8-9 mm long when fully

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-208developed. The head capsule in preservation is light yellow with brown head spots. There is sometimes much infuscation about the head spots (Fig. 91). The gular notch is widest at its base, a little wider than long and is bluntly rounded or broadly pointed at its apex (Fig. 92). The subesophageal ganglion is dark colored, subtriangular in shape. The cephalic fans possess about 41-50 rays and have about 22 primary spines widely separated from each other along the ray with about 12 secondary spines in between each pair of primary spines. The submentum has especially prominent medial and lateral teeth and the margins of the submentum beyond the lateral teeth are conspicuously serrate. There are four to five long and additional short setae on each side of the submentum. The antenna is rather short with two white spots. In alcohol the abdomen is gray or greenish-gray to gray black with lighter intersegmental areas, while in nature the. larva normally appears dark gray black. The anal gills appear as three, thick, simple lobes and anal tubercles are absent or very small. The pupa is located in a slipper-shaued , well-woven cocoon with an anterior edge which slopes back ventral to dorsal and is 3-3.5 mm long. There are 16 filaments in each respiratory organ arranged in 8 pairs which arise at various distances from the base (Fig. 93). The adult male is velvety black with a scutum that bears two anterior bright spots on the sides of a wide medial dark stripe which widens posteriorly and from the posterior sends two curved wide dark projections forward. The distimere is short, stout and bears 3-4 terminal teeth (Fig. 94). The ventral plate in end view is broadly triangu] ar. The wings of the female are 3-3.5 mm long. The female is ash gray

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-209Figure 91. Head spots of a S. vittatum larva. Figure 92. Gular notch of a S. vittatum larva.

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-210Figure 93. S. vittatum pupa and cocoon. {?** Figure 94. Terniinalia of a 5. vittatum male.

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-211Figure 95. Scutum of a female of 5. vittatum. < ..'• >*A Figure 96. Terminalia of a female of S. vittatum.

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-212in color with a silvery gray frons and gray clypeus. The scutum bears 5 conspicuous longitudinal brown marks, the outer pair on each side being almost spots and the middle mark essentially a solid line the length of the scutum (Fig. 95). There is a bold black and gray pattern on the abdomen. The genital fork appears as a widespread-Y with prominent ventral and dorsal projections distally on each arm (Fig. 96). Distribution . In the U.S. S. vittatum has been reported from Alaska (Stone, 1952), Washington (Corredor, 1975), California (Hall, 1974), Utah and Idaho (Twinn, 1938), Kansas (Emery, 1914), Wisconsin (Anderson and Dicke , 1960), Minnesota (Nicholson and Mickel, 1950), New York (Stone and Jamnback, 1955), Delaware (Sutherland and Darsie, 1960a), New Jersey (Crans and McCuiston, 1970), Rhode Island (Dimond and Hart, 1953), Connecticut (Stone, 1964), Pennsylvania (Frost, 1949; Eckhart and Snetsinger, 1969), Massachussetts (Holbrook, 1967), the Tennessee River Valley (Snow et al. , 1958), Maryland (Tarshis, 1968), Virginia (Townsend and Turner, 1976), South Carolina (Garris et al., 1975) and Alabama (Stone and Snoddy, 1969). The latter authors indicate S. vittatum occurs from Alaska and Greenland south to California, Texas, Louisiana, and Georgia and mention that it has not been located below the 29°N latitude with the southernmost records being Uvalde, Texas, and Baton Rouge, Louisiana. I record for the first time its presence in Florida. Life history . Wu (1930) and Stone and Jamnback (1955) mention that S. vittatum females lay approximately 300 eggs in long gelatinous strings on leaves of aquatic plants, stones, and logs in streams . Davies and Peterson (1956) observed females of S. vittatum to fly facing the current of a sluiceway, drop down, touch their abdomens to the water's surface, release eggs, and then fly up and repeat the process. Peterson (1961)

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-213found females oviposit by dipping their abdomens, in flight, to the water of a lake near a waterfall. Holbrook (1967) observed a S. vittatwn female land on a solid object sticking out of the flow, turn, back into the water and lay eggs one inch below the surface. Stone and Snoddy (1969) indicate females will oviposit eggs on almost any substrate at or near the water surface. Wu (1930) found the eggs to hatch in nature at 20-22°C in 4 to 5 days and mentioned that hatching will also occur in standing water but that a longer time is required. Tarshis (1968) found S. vittatwn eggs hatch in two days at 20-21°C in standing water. The eggs of S. vittatwn were found to be unable to resist desiccation (Wu, 1930). The larval stage was found to last 13 to 17 days at 18-26°C (Wu, 1930). Anderson and Dicke (1960) report that S. vittatwn larvae required three weeks for development in the summer in Wisconsin. Tarshis (1965) reported that pupation began 11 to 14 days after larvae were collected in the field. Snow et al. (1958) report the first broods of S. vittatwn in the streams of Tennessee in late February and March. Larvae appear in March in Newfoundland (Lewis and Bennett, 1975). Garris et al. (1975) found larvae and pupae of S. vittatwn only in May in Sumter Co., South Carolina. Wu (1930) found S. vittatwn completed pupation in 3% to 5k days which agrees with the period of 3 to 4 days given by Anderson and Dicke (1960). Tarshis (1965) found adults began to emerge 1 to 2 days after pupation. Wu (1930) reported that a 50:50 ratio of males and females emerged from pupae she observed. Overwintering is reported for S. vittatwn in the egg stage (Lewis and Bennett, 1973), in the larval stage (Dimond and Hart, 1953; Stone and Jamnback, 1955) and in the egg and larval stage (Anderson and Dicke, 1960). Two to three generations a year are reported in Newfoundland (Lewis and Bennect, 1973),

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-214four generations in Rhode Island and New York (Diamond and Hart, 1953; Stone and Jamnback, 1955) and Stone and Snoddy (1969) report at least seven generations per year in Alabama. Ecology . Simulium vittatum is reported to be one of the most abundant and widespread black flies in the irrigation canals of Saskatchewan and Alberta (Fredeen and Shemanchuk, 1960), in the streams and rivers of Wisconsin (Anderson and Dicke, 1960) and everywhere in Alabama except the coastal areas (Stone and Snoddy, 1969). Stone and Jamnback (1955) reported immature S. vittatum populations from dam faces and below lake outlets and large pools. Snow et al. (1958) found immatures on emergent water willow, and vines, grass and weeds trailing in the flow. Wolfe and Peterson (1959) found larvae initially on rocks but observed that they transferred to vegetation when the rocks became covered with algae. Sutherland and Darsie (1960a and b) recovered S. vittatum larvae from rivers and large streams in Delaware. Snow et al. (1958) found S. vittatum to be the most tolerant to silt of all the black fly species they observed and noted that adults would emerge successfully from siltcovered pupae on twigs and rocks. In Wisconsin S. vittatum larvae tolerated large amounts of eroded. soil and other debris in the water courses (Anderson and Dicke, 1960). Immature populations of 2000 to 4000 larvae and pupae per .09 sq m (1 sq ft) were observed in Wisconsin (Anderson and Dicke, 1960). Fredeen and Shemanchuk (1960) report S. vittatum immatures from flows .03-1.5 m/sec (.1 to 5 ft/sec) with temperatures from 6.7°C (44°F) to 32°C (90°F) and pH from 7.3 to 9.6. Stone and Snoddy (1969) report the larvae will tolerate 0°C to 33°C water temperatures, flows from .15 to 1.83 m/sec (.5 to 6 ft/sec), and low pH and oxygen concentrations.

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-215Jamnback (1973) reports transovarial transmission of microsporidian infections in 1 to 4% of surface sterilized eggs of S. vittatum observed in the lab. The fungus Coelomyoidium simulii, often fatal to black fly larvae, was found in S. vittatum larvae in New York (Jamnback, 1973). Fredeen and Shemanchuk (1960) report three sites in Alberta where 95 to 100% of the larvae were parasitized by mermithid nematodes. Phelps and DeFoliart (1964) reported that three genera of mermithids which cause up to 50% larval mortality were found in the larvae of S. vittatum in Wisconsin and they record parasitism of adult S. vittatum by the genera Gastromernris and Isomermis. Molloy and Jamnback (1975) found that the mermithid Neomesomevmis enters S. vittatum larvae by penetrating the integument and achieved, in lab trials, infections in 80% first instar and 64% second instar larvae and 41.9% mortality. Habits . Mulla and Lacey (1976) found that younger S. vittatum larvae fed at a faster rate than older larvae. Bradbury and Bennett (1974a) found black, blue and red were the most attractive colors for S. vittatum adults. Peterson (1961) observed mating swarms of males 1.83-2. 4 m (6-8 ft) above the lip of a waterfall. Hocking (1953) found that S. vittatum flies 258 cm/sec (5.8 mph) and exhibits continuous flight down to a temperature of 12.8°C (55°F) and intermittent flight to 8.9°C (48°F). Davies and Peterson (1956) and Stone and Snoddy (1969) record that males and females of S. vittatum are often collected from blossoms near a breeding site. Adults of S. vittatum were captured in light traps from May to September in Pennsylvania (Frost, 1949). In Alabama S. vittatum was captured in a modified New Jersey light trap with carbon dioxide as the attractant (Snoddy and Hays, 1966). Wu (1930) found that development of the ova in S. vittatum is not

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-216dependent on a blood meal but that the fat bodies provide enough nutrient. Lewis and Bennett (1973) report newly emerged females are autogenous for the first generation. Davies and Peterson (1956) reported that S. vittatum fed on deer in nature and on turkeys and ducks in the lab. Downe and Morrison (1957) found that S. vittatum fed on horses, cattle, and pigs. Snow et al. (1958) report attacks of S. vittatum severe enough to interrupt milking and plowing and drive work stock to cover and record cows being bitten on the udder while with horses and mules most black fly feeding occurred inside the ears. Fredeen (1973) reports that S. vittatum causes severe dermatitis in the ears of horses and cattle in Canada and Townsend and Turner (197 6) report a similar situation with horses in Virginia. Davies et al. (1962) observed S. vittatum feeding on humans on two occasions. Jamnback (1969) indicates S. vittatum is a pest of livestock but rarely attacks man. In Pennsylvania S. vittatum is recorded as a non-biter but a pest of man (Eckhard and Snetsinger, 1969). Lewis and Bennett (1973) indicate S. vittatum is one of the more important pests of man in Newfoundland. Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 2.92 m 14.6 cm 6.1 19.2°C (66.5°F) .54 m/sec (1.76 ft/sec) Min: .075 .5 4.35 10 (50) .14 ( .45) Max: 15.25 90 7.4 27.8 (82) 1.77 (5.8) In Florida 5. vittatum was collected from 44 sites in 21 counties (Fig. 97). Included below is a record of specimens from the Withlacoochee River, Madison Co., which I examined but did not collect. Simulium vittatum is the seventh most widespread black fly in the state

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-217r/ J0 Figure 97. Collection locations for S. vittatum in Florida.

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•218occurring in the wesLern, central and more southern sections. There are five types of flowing water in which the immatures occur: 1) outflows of impounded water such as at spillways (Sites 15, 29, 33, 39,86, 132, 167, 177); 2) sand or mud-bottomed drainage ditches (130, 138, 196); 3) sandy or rocky streams with little vegetation (2, 6, 42, hi , 82, 90, 118); 4) streams clogged with vegetation (34, 72, 162); and 5) spring-fed rivers (181). The largest populations, approaching or exceeding 1000 larvae and pupae per .09 sq m (1 sq ft) were observed below ponds and lakes on spillways usually from January through April (Fig. 98). At Crestvicw (Site 167), the larvae formed a striking black carpet on the orange-colored sediment of the spillway. Larvae and occasionally even pupae appear in October in the streams and S. vittatum is present in most streams until middle or late June (Fig. 8 ). After June perhaps rising water temperatures or growth of unfavorable forms of algae cause S. vittaivjn streams to become unsuitable and populations disappear. In April 1975 a medium to heavy population of S. vittatwn was present at Site 33 on the University of Florida in water with a temperature of 27.2°C (81°F). In July under similar flow conditions and a water temperature of 30°C (86°F) no S. vittatwn immatures were found. Other factors accounting for the disappearance of S. vittatwn could be early summer torrential rains which cause rushing stream conditions and increase the amount of abrasive particulate matter present possibly detrimental to S. vittatum; or, on the other hand, the long sunny, early summer days after the rains result in significant evaporation rates and, often, a decrease in stream flows at some sites, possibly too much to allow S. Vittatwn to remain present. There are no records of S. vittatwn occurring in Florida in July, August, or September although :>est:ivating

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-219Figure 98. Site 167 at Crestview where S. vitiation was collected.

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-220eggs are probably present. Many of the collection records listed below report only small, very young S. vittatum larvae which were found on only one or two of numerous visits to a site. Either the area where the real concentration of this species occurred was missed or, more probably, S. vittatum had difficulty surviving or building up large populations under those particular stream conditions. In sandy streams with sparse vegetation like Holman Branch (Site 82), at Quincy,the water was clouded with silt and only small numbers of S. vittatum along with a few S. tuberosum and S. verecundwn were recovered. At another location, Site 18 a f low to Newnan's Lake, S. vittatum was collected only once in nineteen visits over a period of two years, while six other species were collected there regularly. Perhaps this is an example of competitive exclusion. S. vittatum was found in the drainage ditches mentioned earlier but it was not collected in the main flows at the end of the ditches. The drainage flows were generally more shallow, less wide, warmer, and flowed slower than the main streams. Even in the drainage ditches and definitely in other flows, the largest number of S. vittatum immatures were found where the water channeled or was constricted and increased in velocity such a down a curved concrete ramp (Site 29) , at the entrance to a pipe under the road (33) , or where the stream poured off a concrete bridge support (111). Larvae were collected from the following vegetational substrates in the streams: green filamentous algae, green and brown trailing bank grass, eel grass, dead tree leaves especially those of Magnolia, branches and twigs, soft aquatic plant stems, pine needles, bark, small aquatic bush leaves, roots, the leaves and stems of large elephant ear plants and trailing water willow leaves and branches. In addition larvae were

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-221found on rocks, occasionally, and concrete, cardboard, black plastic, firm but gritty sand, and a wooden stake. Pupae were attached to many of the same substrates especially twigs, dead tree leaves, grass, green filamentous algae, and pebbles. The summary of stream parameters above reveals that the immatures of S. vittatwn were usually found in small to medium-sized streams with a moderate to swift flow that exhibited a slightly acidic to neutral pH reaction. The species association table (Table 3 ) reveals that S. vittatwn was collected with ten other species in the streams of Florida. Most frequently S. tuberosum and S. vereoundwn were cohabitors with S. vittatwn. Much less often S. slossonae, S. lakei and S. jonesi were found with S. vittatvm. S. decovwn, which often is found at the outflows of impounded water as is S. vittatwn, had the advantage over S. vittatwn at Shepard's Mill (Site 86) where S. vittatwn was only collected once and S. deaorwn was found many times. In all other outflow situations S. vittatwn populations greatly exceeded those of S. decorwn. Only on one occasion during mid-March at the Crestview spillway (Site 167) were the numbers of S. dezovwn observed to approach those of S. vittatwn. In a few collections unusual forms of S. vittatwn have been noted. In Hogtown Creek (Site 15) a white S. vittatwn larva was collected but was judged to have just molted. Further downstream (Site 32) a group of S. vittatwn larvae and some of S. verecundum were found all of which were whitish in color. The S. vittatwn larvae appeared to have normal characteristics except for being unpigmented. Subsequent collections from the same site yielded slightly darker but still whitish gray S. vittatwn larvae. Pupae have been found with 14 rather than 16 filaments in one of the respiratory organs and one pupa examined had respiratory organs

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•222with 14 and 15 filaments. Only on two occasions were larvae observed with microsporidian infections. No mermithids have been noted in S. vittatwn larvae. Four adult female S. vittatwn were captured during early May in a carbon dioxide-baited Manitoba trap near a stream containing S. vittatum immatures (Site 34). During March at the Tall Timbers Research Station in Leon County the ovipositing behavior of S. vittatwn was observed. A silvery S. vittatwn female, which was later captured and placed in alcohol, was observed at about 1700 hours to tap her abdomen while in flight to water pouring down a 45° concrete incline. The 1.2 m (4 ft) long concrete incline handled overflow from a large open swamp. The water on the spillway was 1-2 cm deep and flowed .61 m/sec (2 ft/sec). The female dipped down and touched the water two or three times as she flew up the incline. Other females flew very close to the water just above the lip of the spillway and may have been dropping eggs there also. Florid a collection records for S. vittatum . Alachua Co. 1)1974: 7 Marcb (S) . 2)1974: 25 May (M) . 7)1974: 2 Feb (S), 7 March (S) , 16 May (M,P) ; 1975: 12 Feb (S), 8 April (S). 11) 1973: 2 Nov (S) . 13) 1974: 4 May (S) . 15) 1973: 7 Dec (S,M,P); 1974: 2 Jan (S,M), 9 March (S,M,P,C), 13 April (S,M,P), 9 May (S,M,L,P,C), 7 June (S) , 19 Oct (S,M), 16 Nov (S) , 14 Dec (S,M,P); 1975: 31 Jan (S,M), 4 April (S,M,L,P,A), 13 May (S). 16) 1976: 1 3 March (S) , 20 March (S,C). 18) 1975: 10 Jan (M). 22) 1974: 7 Dec (S) . 29) 1974: 13 April (S) , 9 May (S). 32) 1974: 2 Feb (S,P), 9 March (3,L). 13 April (P) , 16 Nov (L) ; 1975: 10 Jan (S,M,P,A), 4 April (M) . 3.3) 1974: 9 Jan (S.M.L.P), 14 Feb (S.M.C), 10 April (S.M.L.P), 18 May (S.M.L.P.A); 1975: 8 Jan

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-223(S,M,L,P), 1 March (S,M,L,P,A), 30 April (S,M,L,P,A), 18 June (S,C), 22 Oct (P); 1976: 5 April (S,M,L,P,C,A) . 34) 1974: 9 Jan (P,A), 13 March (S,M,P,C), 10 April (S,M,P), 3 May (A), 16 May (S,M,P); 1975: 8 Jan (S,M,P,C), 1 March (S,M,L,C), 30 April (S,M,L,P,C), 18 June (S,M,L,P); 1976: 5 April (S,M,L,P,A). 39) 1974: 2 Jan (S,M,L), 12 Jan (S) , 14 Feb (S,M), 2 April (S,M,L,P,C), 16 May (S,M,L,P), 30 June (M) , 25 Oct (S,M,L,P,A), 23 Nov (S,M,L,P,C,A) ; 1975: 10 Jan (S,M,L,), 1 March (S,M,L), 30 April (S,M,L,P,A), 19 Nov (S). Bradford Co. 42) 1975: 5 April (S). 47) 1975: 26 April (S). Dixie Co. 68) 1975: 17 Jan (S,L). Duval Co. 72) 1974: 20 April (S,M); 1975: 12 March (S.M.L.P), 4 May (S,M). Gadsden Co. 81) 1974: 17 March (S) ; 1975: 27 March (S). 82) 1974: 17 March (S.M.L.P); 1975: 18 Jan (S,L,P), 27 March (S) , 10 June (S,M,P). 83) 1974: 17 March (S,M,L,P,C); 1975: 18 Jan (S,M,L,P,A), 27 March (S,M,L,P,C,A) . 86) 1975: 18 Jan (P,A). 87) 1975: 27 March (S). 90) 1974: 17 March (S); 1975: 18 Jan (S.M.L), 27 March (S), 10 June (S), 29 Dec (S,M,L). Hamilton Co. 100 ) 1976: 24 Jan (M) . 101 ) 1975: 1 Feb (L) . Hardee Co. 102 ) 1975: 22 March (S,M,L,P). Hillsborough Co. Ill ) 1975: 22 March (S,M,L,P,C,A) . Holmes Co. 118 ) 1975: 28 March (S,M,L,P,A), 11 June (S,M,L,C). Lafayette Co. 130 ) 1974: 14 June (S,M,L,P,A); 1975: 26 March (S). Leon Co. 132 ) 1974: 16 March (S,M,L,P,C,A) . Levy Co. 138 ) 1975: 31 Jan ( M) . Madison Co. 152 ) 1967: 30 Nov (S,M,L W. Beck).

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-224Nassau Co. 162 ) 1974: 20 April (S); 1975: 12 March (P) . Okaloosa Co. 167 ) 1974: 18 March (S,M,L), 16 June (S,M,L,P), 13 Oct (S,M,L,P); 1975: 19 Jan (S,M,L,P,A), 28 March (S,M,L,P,A), 31 Dec (S); 1976: 18 April (S,M,L,P). Pasco Co. 177) 1975: 23 March (S,C). Polk Co. 181) 1975: 22 March (S). 183 ) 1975: 22 March (S) . Putnam Co. 186 ) 1975: 12 Feb (S). Santa Rosa Co. 196 ) 1974: 18 March (S). Seminole Co. 202 ) 1974: 21 March (L) , 12 May (S,M,P), 28 Nov (S) ; 1975: 15 March (S,L,P), 27 April (S,M,L,P,C). 203 ) 1975: 15 March (S,P), 27 April (S,P). Taylor Co. 210) 1973: 17 Dec (S,L). 213 ) 1975: 17 Jan (S). Simulium (Simulium) decorum Walker Simulium decorum Walker, 1848, List Dipt. Brit. Mus. 1 : 112 (adult). Simulium pisaioidium Riley, 1870, Amer. Entomol. and Bot. 2: 367 (larva and pupa only) . Simulium venustoides Hart, 1912, in Forbes, 111. State Entomol Rep. 27: 42 (female only). Simulium decorum katmai Dyar and Shannon, 1927, Proc. U.S. Nat. Mus. 69(10): 31 (female). Simulium ottawaense Twinn, 1936, Can. J. Res., D, 14: 146 (female, male, pupa) . Simulium decorum — Stone, 1964, Conn. State Geol. and Natur. Hist. Surv. Bull. 97: 44 (female, male, larva, pupa).

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-225Taxonomy . Walker (1848) described an adult, apparently a female, from St. Martin's Falls, Albany River, Ontario. The holotype female is located in the British Museum, London, England (Davies et al. , 1962). Stone (1964) mentioned that the relationship between S. decorum and supposedly synonymous European species like S. noelleri was still unclear. Stone and Snoddy (1969) suggest S. argyreatum Meigen, with which 5. noelleriFrederichs is now considered synonymized, may be appropriate for the far north form in America which Dyar and Shannon called S. decorum katmai. Description . The larva is 7.5-8 mm long, is gray in appearance in the preserved state and blackish gray in nature. The head capsule is a combination of light and dark brown with a dark brown fulvous area longitudinally on each side of a prominant median row of white head spots (Fig. 99). The cephalic fans each consist of about 57 rays. The gular notch is light colored, arrowhead-shaped, pointed distally, and extends about one half the distance to the teeth of the submentum (Fig. 100). The pupa is located in a rather loose, coarse, cocoon 4-4.5 mm long and bears 8 thin filaments on each respiratory organ (Fig. 101). The male wings are 2.5-3 mm long. The scutum is velvety black with many thin golden hairs and bears a faint silvery spot at each anterior lateral angle. The distimere is three times longer than wide and about twice as long as the basimere. The ventral plate in end view bears a median portion which is narrow and elongate (Fig. 102) and in lateral view displays a ventral keel. The female wings are about 3 mm long. The base of the radius is bare. The frons and abdominal tergites are gray pollinose. The antennal

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-226Figure 99. Head spots of a S. decorum larva. *+ Figure 100. Gular notch of a 5. deaonan larva.

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-227Figure 101. S. decorum pupa and cocoon. Figure 102. Male terminalia of S. decorum.

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-228Figure 103. Female terminalia of S. decorum.

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-229scape and pedicel are orange brown and the flagellum is dark brown. The fore tibia bear a dorsal elongate white patch. The legs are yellow brown with the segments darkening distally. The scutum is dark brownish gray in color and convex in lateral view. The genital fork is in the form of a broad Y with tapering arms that end in blunt, prominent, subapical projections (Fig. 103). Distribution . Shewell (1957) lists S. decorum as Nearctic and widespread in northern America. Stone (1964) mentions this species is found from Alaska to Newfoundland and south to Oregon, Colorado, Arkansas, and Florida. Jones and Richey (1956) collected it in South Carolina, Snow et al. (1958) report it in their survey of the Tennessee River Valley and Stone and Snoddy (1969) found it commonly in central and northern Alabama. Life history . In Ontario eggs overwinter and hatch during April (Davies et al., 1962). DeFoliart (1951) found the eggs hatch within seven days at 21.1°C (71°F). Tarshis (1968) found S. decorum eggs hatch in two days in standing water at room temperature (20-21°C) . Stone and Snoddy (1969) report overwintering occurs as eggs, larvae or pupae. Anderson and Dicke (1960) mention first generation S. decorum larvae take 4 weeks to mature, second generation larvae take 3 weeks and pupae develop in 3 to 5 days. Davies et al. (1962) state that pupation occurs in mid-May and adult emergence takes place in late May. Abdelnur (1968) observed at least three cycles of pupation during 16-20 June, 22-25 July, and 11-19 August. Jones and Richey (1956) observed overlapping generations in South Carolina from February to May and suggest that with sufficient rain S. decorum breeds all summer with a generation every month. Stone and Snoddy (1969) indicate the initial mass emergence of adults

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-230occurs during late April and a new generation is completed every five or six weeks thereafter until early November. Ecology . Stone and Jamnback (1955) state that S. decorum larvae almost invariably occur on dams, at lake outlets or below large pools. 2 2 Jones and Richey (1956) found 442 larvae per 6.45 cm (1 in ) on a dam spillway in April and Anderson and Dicke (1960) report populations of 2 2 1000-2000 larvae per .093 m (1 ft ) of substrate. Davies et al. (1962) found larvae in streams less than . 3 m (1 ft) to over 4.6 m (15 ft) wide. Stone and Snoddy (1969) found S. decorum in streams with temperatures ranging from 0° to 33°C and currents up to 1.83 m/sec (6 ft/sec). Hudson and Hays (1975) found that in trough tests S. decorum larvae preferentially attached to substrates with particles in the .07 to .1 mm size range rather than substrates with smaller or larger particles. Larvae were found more abundantly in areas of dense plant parts such as the inner surfaces of root clusters. Ezenwa (1973) reported mermithid (Neomesomermis) and microsporidian (Th.eloha.nia and Pleistophora) infections in S. decorum. McKague and Wood (1974) found a granular SRIO Altosid formulation at .1 ppm concentration was 100% effective in halting adult emergence. Habits . Females oviposit eggs in irregular masses while settled on vegetation, wood, concrete or rocks at the outlets of impounded water where a thin film of water flows over the substrate or, in more protected locations, may tap the surface of the water and drop one or more eggs into the water while in flight (Davies and Peterson, 1956; Davies et al. , 1962). Adults may mate on the ground soon after emergence with the male approaching the female ; crawling on her back, and curving his abdomen down to touch the tip of the female's abdomen (Davies and

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-231Peterson, 1956). Davies and Peterson (1956) reported that S. decorum occasionally fed on humans, and in vials inverted over bare skin it fed on ducks, geese, crows and turkeys. Stone (1964) mentions this species is frequently a serious pest of horses, cattle and deer and suggest that it may attack birds. Jamnback (1969) lists S. decorum as mammalophilic but rarely a man-biter and mentions adults were found harboring developing trypanosomal stages. Stone and Snoddy (1969) found in Alabama that it attacks cattle and regularly bites man but was not considered a serious pest of humans. Florida observations . Stream Parameters Width Depth pH Temperature Mean: 7.37 m 7.7 cm 4.96 18.8°C (65.8°F) Min: .1 .5 4.25 8.9 (48) Max: 36 60 7.0 26.7 (80) Velocity .82 m/sec (2.7 ft/sec) .3 (1) 1.83 (6) Simulium decorum is recorded from 11 scattered locations in 5 Florida counties (Fig. 104). In almost every case the flow in which immatures of this species were found was immediately below or a short way downstream from a body of impounded water. A more specific search for black flies at locations such as on dams and at the outlets of lakes would most likely reveal a much wider or more uniform distribution for this species in Florida. At Shepard's Mill (Site 86), Chattachoochee (88) , and Crestview (167) S. decorum was found on concrete spillways or dams where the flow reached 1.83 m/sec (6 ft/sec) at times. The width of the flows where S. decorum was found varied from 1 m or less at Site 33 on the Univ. of Florida campus and Site 132 at Tall Timbers to 12 ana 36 m on the concrete at Shepard's Mill and the Chattahoochee Dam,

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-232rr _«^ ^fit. -^ Figure 104. Collection locations for 5. decorum in Florida,

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-233respectively. The depth of the water at the collection sites was usually less than 10 cm. Larvae and pupae were found attached to pine needles, trapped grass, dead leaves, algae, concrete and orange-brown sediment. Collections at most locations were made from late December through mid June. At Shepard's Mill (Fig. 105) S. decorum was present in all stages all year. Water poured from Shepard's Pond over a series of 1.8 m wide and about 2 m high concrete and wood spillways next to each other onto a gently sloped, algae-covered, concrete run, passed under the road through three culverts and emptied into a large pool below. Heavy populations of larvae and pupae were found, especially during March, concentrated and lined up in narrow bands along irregularities in the concrete, more evenly distributed in large mats on the concrete and entangled in long thin green algae where the flow was a few centimeters deep and .3-1.5 m/sec (1-5 ft/sec) or faster. At this site during March females were observed flying back and forth at the crests of the spillways tapping their abdomens, presumably ovipositing, to the smooth water just before it plunged toward the concrete. Other females of S. decorum were removed during March and June from spider webs constructed on the supports of a few of the less active spillways. Reflecting its presently known restricted distribution and preferred habitat S. decorum was associated with only five other black fly species in Florida's flows (Table 3 ). Simulium vittatwn and S. verecundum were associated on nine and seven occasions, respectively. Simulium tuberosum, S. jonesi, and S. noziale were collected with S. decorum on six, three, and two occasions, respectively. Simulium decorum females were identified as the flies involved in two human biting incidents, both in Okaloosa County: on 22 March in

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-234Figure 105. Shepard's Mill, Site 86, where S. decorum was collected,

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-235Crestview and 28 May in Destin. Female adults were captured on the wing in late March in Jefferson County and during early April in Alachua County. Florida collection records for S. decorum . Alachua Co. 33) 1976: 5 April (S) . 34) 1976: 6 April (A). Gadsden Co. 83) 1975: 27 March (M) . 86) 1974: 17 March (S,M,L,P,A), 6 Aug (S,M,L,P,C), 13 Oct (S,M,L,P,C); 1975: 18 Jan (S,M,L,P,C,A) , 27 March (S,M,L,P,C,A) , 11 June (S,M,L,P,C,A) , 24 Aug (S,M,L,P,C), 29 Dec (S,M,L,P,C,A). 88) 1975: 30 Dec (S,L); 1976: 17 April (S,M,L,P,A). 90) 1975: 18 Jan (S,M,L), 27 March (S,M,L,C). 91) 1975: 27 March (L) . Jefferson Co. 125 ) 1975: 26 March (A). Leon Co. 132 ) 1974: 16 March (S,L,P,C). Okaloosa Co. 167 ) 1974: 18 March (S,M,L), 16 June (S) ; 1975: 12 June (S,M,L); 1976: 22 March (A), 18 April (S) . 224) 1976: 28 May (A). Simulium (Simulium) tuberosum (Lundstrom) Melusina tuberosa Lundstrom, 1911, Acta Soc. pro Fauna Flora Fenn. 34(12): 14 (male). Simulium peris sum Dyar and Shannon, 1927, Proc. U.S. Nat. Mus. 69(10): 43 (female, male) . Simulium vandaliaum Dyar and Shannon, 1927, Proc. U.S. Nat. Mus. 69(10): 44 (male) . Simulium turmale Twinn, 1938, Can. Entomol. 70: 51 (male). Simulium twinni Stains and Knowlton, 1940, Ann. Entomol. Soc. Amer. 33: 77 (male).

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-236Simulium tuberosum, Stone and Jamnback, 1955, N.Y. State Mus. Bull. 349: 78 (female, male, pupa, larva). Simulium tuberosum, Stone, 1964, Conn State. Geol. Natur. Hist. Surv. Bull. 97: 45 (female, male, larva, pupa). Taxonomy . Lundstrom in 1911 in Finland first described a male of this species now known as Simulium tuberosum. Stone (1964) indicates that the syntypes are probably in the Kansallismuseo in Helsinki. Stone and Jamnback (1955) in their work on the black flies of New York State included descriptions of larva, pupa, and adults of S. tuberosum. Davies et al. (1962) and Jamnback (1969) suggest that S. tuberosum is a species complex of two or more undescribed forms. Landau (1962) separated four sibling forms of S. tuberosum based on variations in their salivary gland chromosomes. Stone and Snoddy (1969) mention two ecologically different forms in Alabama and suggest that if S. tuberosum in Europe is determined to be different from the form in North America, S. perissum Dyar and Shannon would be the most correct name for the southeastern United States species. Description . Last instar larvae of S. tuberosum are 5. 5-7. mm long. The abdomen is dark steel blue gray or reddish brown in color. The cephalic fans contain 37-42 rays. The head spots are darkish but indistinct and often appear as part of a fulvous pattern on the posterior portion of the cephalic apotome (Fig. 106). The gular notch is longer than wide, extends about one-half the distance to the submental teeth and is pointed or narrowly rounded at the apex (Fig. 107). The subesophageal ganglion is dark. The anal tubercles are small and inconspicuous.

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-237Figure 106. The cephalic apotome of a S. tuberosum larva. Figure 107. The gular notch of a S. tuberosum larva.

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-238Figure 108. Pupa and cocoon of S. tuberosum. * Figure 109. Male terminalia of S. tuberosum.

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-239v?aft Figure 110. Terminalia of a female of S. tuberosum.

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-240The pupa is 2.5-3 mm long in a fairly uniformly textured, lightbrown, slipper-shaped cocoon. The respiratory organ consists of six thin filaments about one-half the length of the pupa which rise in pairs from three short petioles (Fig. 108). The dorsal filaments are thicker than the ventral filaments. All pairs of filaments are fairly equally spaced from each other at the bases. The males are velvety black in appearance with an oblique, silvery shiny patch at both antero-lateral angles of the scutum. Each curved, elongate distimere bears a number of spines on its basal lobe and the ventral plate in ventral view appears as in Fig. 109. The female bears wings 2.5-3 mm long. The female is dark, gray and shiny in appearance with a shiny black frons and has hairs under the subcosta. Females possess a long dull white patch on each tibia which is not wider than one-third the width of the tibia. The ovipositor flaps or lobes are fairly parallel sided. The tarsal claws lack a basal tooth. The terminalia are as in Fig. 110. Distribution . Shewell (1956) and Stone and Snoddy (1969) list S. tuberosum as circumboreal and holarctic, respectively. Davies et al. (1962) indicate that the type locality is probably Enontekis, Finnish Lapland, Finland. Ussova (1961) reported S. tuberosum from the U.S.S.R. while Grenier (1953) recorded it in France and Edwards (1915) and Smart (1944) noted its presence in England. Stone (1952) found S. tuberosum in Alaska. Stone (1964) lists the distribution of this species in the Nearctic region from Alaska and Greenland south to California, Texas and Florida. Life history . Davies et al. (1962) indicate that in Canada the eggs overwinter and that there are three generations each year. Holbrook

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-241(1967) found S. tuberosum to overwinter as eggs and to be multivoltine in Massachusetts. In Alabama overwintering occurs in the larval, pupal and adult stage (Stone and Snoddy, 1969). Davies et al. (1962) found that females emerge with undeveloped eggs and a moderate amount of stored nutrient. In Newfoundland the first generation larvae are present three to four weeks, the pupal stage lasts four to six days, the first generation adults emerge during mid-May and egg hatching of the second generation occurs two months later (Lewis and Bennett, 1973). Five or more generations are completed each year in Alabama (Stone and Snoddy, 1969). Ecology . Sommerman et al. (1955) reported heavy (1500 larvae/ .09 sq m (1 sq ft)) populations of S. tuberosum in Alaskan streams and observed parasitism by mermithids in up to 65% of the individuals in some streams. Garris and Noblet (1975) report mermithid parasitism in South Carolina in S. tuberosum larvae from January through December ranged from 5 to 40%. Snow et al. (1958) found that S. tuberosum was the most frequently encountered black fly in the Tennessee Valley occurring in large numbers especially in mountain streams and in shallow, gravelly, well-exposed flows in lower areas. Ussova (1961) reported S. tuberosum from mountain rivers .5-2 m deep and found immatures attached to rocks and aquatic vegetation where the current was .5-1.2 m/sec in large and small creeks originating from lakes. Turbid, polluted water with an oxygen content less than 55% was rarely inhabited by S. tuberosum. Lewis and Bennett (1973) observed no substrate preference in S. tuberosum larvae and collected larvae from streams .5-20 cm deep with velocities from .27 to 1.34 m/sec and temperatures 6-18. 5°C. Habits. Davies and Peterson (1956) failed to observe S. tuberosum

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-242females laying eggs on solid substrates and suggest that this species drops eggs freely into the water while flying. Wolfe and Peterson (1959) observed females of S. tuberosum land on the calm surface of stream water between stones and deposit about twenty eggs which sank to the bottom. Downe and Morrison (1957) reported that S. tuberosum obtained blood from horses, cows, and chickens in a barn. Snow et al. (1958) collected S. tuberosum while it was feeding in the ears of horses and mules and crawling upon but not biting men about the head. Stone and Snoddy (1969) state that S. tuberosum is a serious pest of man and livestock in Alabama, feeding on the ears, udder, abdomen, and genitalia of cattle and readily biting people engaged in outdoor activities. Florida observations . 1 Stream Parameters J Width Depth pH Temperature Mean: 2.24 m 22.93 cm 4.69 17.4°C (66.3°F) Min: .03 1 3.5 7.2 (45) Max: 10 100 7.6 29.4 (85) Velocity .52 m/sec (1.7 ft/sec) .153 ( .5) 1.53 (5) Simulium tuberosum was collected from 97 locations in 30 counties in Florida (Fig. ill). This species is the second most abundant black fly in the state and is present in some streams all year round. The distribution of S. tuberosum includes locations in Escambia County in west Florida, points all across the coastal plains in the northern part The stream parameters above are compiled from figures gathered at over 50 sites involving more than 330 collections where at least three of the stages S, M, L, or P of S. tuberosum were found. Parameters from sites where S. tuberosum was less well represented, such as the big rivers, are not included.

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-243Figure 111. Collection locations for S. tuberosum in Florid;

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-244of the state, generally high ground in the north central area and records as far south as Highlands County and Hardee County just northwest of Lake Okeechobee. This species was not collected in many counties bordering the Gulf of Mexico although a considerable number of collecting attempts were made for example in Levy County and Taylor County. In the counties of Alachua, Clay, Gadsden and Liberty S. tuberosum was found in many of the streams checked. Simulium tuberosum was found occasionally in large rivers such as the Santa Fe (Site 58), the Blackwater (1G3) , the Withlacoochee (151) and the Suwanee (208) however only in small numbers. Occasionally, in the spring, in the shallow flows of concrete drainage ditches near larger main streams such as at the Patuxent Research Station (Site 39) and at South Prong Black Creek (57) S. tuberosum was the only species present and dense populations were encountered. There are two general types of streams, however, where S. tuberosum was usually present all year and reached its greatest numbers: small, sandy flows with little vegetation and small flows with considerable vegetation. The first category involves streams where S. tuberosum was usually the sole or at least the predominating species present and reached moderate to heavy numbers. These streams include Blues Creek (Site 2), a stream at the Devil's Millhopper (3), Gold Head Branch (55), Rock Creek at Torreya State Park (148) and Adam's Mill Creek (220) . In these shallow, 1-3 m wide flows the substrate was sand or pebbles and sand and immature*, were found attached to pine needles, dead leaves, roots, pebbles, rocks, clay and, occasional ly, some trailing grass, sedges or other green aquatic vegetation. • Most of these flows occurred in wooded, shaded situations.

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-245The largest populations of S. tuberosum were discovered in streams of the second category with much vegetation such as the small flow at Site 8, Hatchet Creek (17), Site 56 (Fig. 112), West Minnow Branch (144), Thomas Creek (159) and Palmetto Branch (187). These streams were normally not more than 1 m wide or 30 cm deep. Abundant trailing grass, or large leafed and small leafed aquatic vegetation was located in the flow and it was on this flora plus trapped dead leaves that heavy numbers of S. tuberosum were found. Another favored habitat of S. tuberosum, was en the concrete of highway culverts where the stream poured off to continue its journey below. Good populations of immature S. tuberosum were found on the concrete, algae, leaves and other trapped debris in such situations at the tributary to Telogia Creek (Site 142) and Muddy Brook (Site 165). As typical of most black fly species the immatures of S. tuberosum were usually found concentrated where the flow was the fastest. Site 148 at Torreya State Park illustrates this with the largest populations being found attached to the rock-hard surface of clay boulders over which a thin film of water poured down a meter high falls. At Site 8 during February 1974 large numbers of S. tuberosum were found concentrated in a 15 cm wide, 6 cm deep, grassy channel. At Palmetto Branch (Site. 187) pools above and below the collection site restricted the current and black fly immatures to a sloped section a few meters long. In rushing Big Juniper Creek (Site 195) S. tuberosum was collected from algae-bearing rock-hard clay substrate, twigs and trailing water oak leaves where the water velocity often was 1.22 m/sec (4 ft/sec). The stream populations of 6". tuberosum were usually at their lowest during mW summer, began to increase by October, and reached maximum

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-246Figure 112. Site 56 in Clay County, a collect! on spot for S. tuberosum.

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-247numbers from January through May. This species is multivoltine in Florida and the seven or more generations each year overlap considerably. Since S. tuberosum has adapted to a wide array of streams and is present all year long it was found in association with a great number of other black fly species. Fifteen other simuliid species were collected with S. tuberosum in Florida's streams and rivers (Table 3 ). Most frequently S. tuberosum was found with S. jonesi and S. slossonae , often on the same leaf, pine needle or other substrate. Simulium lakei and S. oongareenarum were collected next most commonly with S. tuberosum. Simulium tuberosum larvae were observed with retarded histoblast growth and bodies swollen with large white uniform masses or filled with many tiny white spheres which were interpreted as protozoan infections. Such infections were encountered in small larvae at six sites and, in larger populations, the infection rates ranged from 2.3% to 8.6%. Mermithids were found in S. tuberosum larvae at ten locations and were observed in large or well-developed larvae once. Parasitism rates ranged from 1.3% (1 of 77 small larvae) to 39% (16 of 41) with most parasitism occurring in less than 10% of the small larvae collected. At one site (124) in Jefferson County mermithids were found parasitizing S. tuberosum larvae on 6 of 7 visits which covered all seasons of the year. It was at this site that 2 of 11 large larvae collected contained mermithid nematodes in the abdominal region. The respiratory histoblasts of these larvae were dark, well-formed, but about one-half normal size. Two color variations in S. tuberosum larvae a reddish-brown form and a steel-blue form have been observed occurring alone in some streams and together at other sites. The gular notch appears more angular and pointed in the gray-blue form and more rounded apically in

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-248the brown form. Variations in the ventral plate of the male have been noted for example whether the arms in ventral view are straight or curve outward and differences in the width of the main lower portion of the plate. From Site 102, Troublesome Creek, in Hardee County a male was reared which possessed a typical ventral plate but had claspers more broad than normal for S. tuberosum. Simulium tuberosum in Florida may actually be a species complex. Adult females were captured in Manitoba traps (Table 4 ) and Malaise traps (Sites 2 and 25) during the spring. Reared females of S. tuberosum were given the opportunity to feed on turkeys in the laboratory on eight occasions from August to January. None of the females engorged on the birds. Florida col le ction records for S. tuberosum . Alachua Co. 1) 1973: 7 Dec (M) , 13 Dec (S,P); 1974: 2 Feb (S,M,L,P), 7 March (S,M,L,P,A), 12 April (S,M,L,P), 6 July (S) , 20 July (S) ; 1975: 10 Jan (S,M,L,P,C,A) , 12 Feb (S,M,L,P,C,A) , 4 April (S,M,L, P,C,A), 10 May (P,C), 15 July (L) , 9 Nov (S) ; 1976: 21 Jan (S,M,L,P,A) 2) 1973: 25 Aug (S,M,L,P), 11 Oct (M,P,A); 1974: 2 Jan (S,M), 14 Feb (S,M,L,P), 2 April (S,M,L,P), 25 May (S,M,L,P), 30 June (S,M,L,P,C), 17 Aug (S,M,L,P), 21 Sept (S,M,P,C), 14 Dec (S) ; 1975: 10 Jan (S,M,L,P,A), 1 March (S,M,L,P,C), 19-20 and 21-23 March (AG.B. Fairchild), 30 April (S,M,L,P,C), 26 May (P,C), 3 Oct (S,M,L,P,C,A), 10 Dec (S,M,L,P,C); 1976: 9-10 March (AJ. Click). 3) 1973: 25 Aug (S,M,L); 1974: 2 April (S,M,P,A), 7 June (S,M, L,P), 14 Sept (S,M,L,P,C), 30 Oct (S,M,L,P,C); 1975: 10 Jan (S,M, L), 26 May (S,M,L,P), 15 July (S,M,L), 3 Oct (S,M,L,P,A). 6) 1973: 22 Sept (S,M,L,P,A), 16 Oct (S,M,L,P), 25 Oct (S,M,L,P); 1974:

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-2492 Jan (S.M.L.P), 2 Feb (S,M,L,P), 7 March (S.M.L.P), 2 April (S,M,L,P), 18 May (S,M,L,P,C), 30 June (S.M.L.P.C), 30 July (S,M), 14 Sept (S.M.L.P.C), 10 Oct (S,M,P);1975: 15 Jan (S,M,L,P,C,A) , 5 April (S,M,L,P,C,A), 18 May (S,M,L,P,C), 1 July (S,M,C), 16 Aug (S,M,L,C), 8 Oct (S,M,L,P,C), 6 Dec (S,M,L,P,C). 7) 1973: 13 Dec (S,M,L,P,C); 1974: 2 Feb (S.M.L.P), 7 March (S,M,L,P), 16 Nov (S,M,L,P,A), 14 Dec (S,P). 8) 1973: 5 Sept (S,M,L), 2 Nov (S,M, L,P); 1974: 14 Feb (S,M,L,P,C), 2 April (S.M.L.P), 13 May (S.M.C), 30 June (S.M.L.P), 3 Aug (S,M,L,P), 22 Sept (S.M.P), 25 Oct (S,M, L,P,A); 1975: 10 Jan (S,M,L,P), 12 Feb (S,M,L,P,C,A) , 4 April (S,M,L,P,C,A), 18 May (P,C), 1 June (S.M.P.C.A), 18 June (S.M.L.P), 15 July (S,L), 25 July (P) , 31 July (S,M,L,P,A), 30 Aug (S,C), 30 Sept (S,M,L,P,C), 9 Nov (S,M,L,P,C), 14 Dec (S,M,L,P,C,A) ; 1976: 6 March (S,M,L,P,C). 11) 1973: 2Nov(S). 13) 1975: 8 Jan (S,M), 30 April (C). 15) 1973: 7 Dec (M,L); 1974: 2 Jan (S) , 9 March (S,M), 16 Nov (S,M,L,P), 14 Dec (S,M,P). 16) 1976: 13 March (S,M,L,P), 20 March (S,M,L,P,C). 17) 1974: 2 Jan (S,M,L,P), 2 Feb (S,M,L,P,C), 7 March (S,M,L,P,A), 12 April (S,M,L,P,C), 5 May (S,M,L,P), 6 June (S,M,P,A), 1 July (S,M,L,P), 4 Dec (S) ; 1975: 11 Jan (S,P,C), 1 March (S ,M,L,P,C,A) , 30 April (S,M,L,P, C,A), 21 June (C) , 16 Aug (S) , 6 Dec (S,M,L,P). 18) 1974 : 12 j an (S,M,L,P 5 C,A), 2 March (S,M,L,P,C,A) , 11 April (S,M,L,P,A), 25 May (S,M,L,P), 30 June (S,M,L,P,C), 24 July (M,L,P,C), 24 Aug (S,M,L,P), 22 Sept (M,L,P), 25 Oct (S,M,L,P), 23 Nov (S,M,L,P); 1975: 10 Jan (S,M,L,P,C,A), 31 Jan (S,M,L,P,C,A) , 8 April (S ,M,L,P,C,A) , 11 May (S,M,L,P,A), 14 June (S,M,L,P,C), 17 July (S,M,L,P,C), 1 Sept (S,P,A), 17 Oct (S,P), 10 Dec (S,M,L,P,C,A) . 20) 1974: 12 Jan

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-250(P), 2 March (M,L), 12 April (P,C); 1975: 31 Jan (S), 18 April (S,M). 21) 1974: 2 March (S,M); 1975: 24 Jan (S,M), 4 April (S,M); 1976: 29 Jan (P) . 22) 1974: 7 Dec (S) ; 1975: 24 Jan (S,L); 1976: 6 March (S) . 24) 1976: 19 Feb (P) . 25) 1975: 13 March and 15 March (AJ. Glick) . 26) 1974: 13 April (S,M,L) 18 May (S,M,P,C). 27) 1974: 24 July (S,C). 28) 1975: 28 Oct (S), 9 Nov (S,M,L,P,C), 10 Dec (C) ; 1976: 30 Jan (S) , 26 March (A). 32) 1974: 9 March (L,P) , 16 Nov (S) ; 1975: 10 Jan (S C,A); 1976: 21 Jan (S,M,L,P,A). 33) 197 L.P. 4: 9 Jan (P); 1975: 28 Nov (S). 34) 1974: 9 Jan (S); 1975: 28 Nov (S,M). 36) 1974: 12 April (S,M,L), 6 July (S,M); 1975: 15 Ian (S,M,L,P), 4 April (S,M,L,P,C,A), 22 May (S), 22 Oct (S,M,L,P,C,A) . 39) 1974: 12 Jan (S,M), 14 Feb (P,C), 2 April (S) , 25 Oct (S,M,P), 23 Nov (S,M,L,P); 1975: 10 Jan (S.M.L.P), 1 March (S,M), 30 April (S,M,L,P,C,A) , 26 July (S.M.L.P), 27 Sept (S), 19 Nov (S,M,L,P,A); 1976: 21 Jan (S,M,L,P,C). Bay Co. 40) 1975: 27 March (S,M,L,P), 11 June (S,M,L,P,C) , " 6 Sept (S,M,P,A), 30 Dec (S.P.C.A). Baker Co. 41) 1974: 4 Jan (S,M,L,P), 1 June (S.M.L.P), 1 7 Sept (S,M, L,P), 6 Nov (S,M,L,P,C); 1975: 1 Feb (S,M,L,P,C,A) , 26 April (S.M.P), 21 June (S.M.C), 17 Aug (S,M,L,P,C), 14 Nov (S,M,L,P) Bradford Co. 42) 1973: 6 Oct (S,L,P); 1975: 19 Nov (S,M,L,P,A) 43) 1975: 5 April (P); 1976: 7 Jan (S) , 21 Jan (S,M). 45) 1975 26 Jan (S) , 5 April (S). 47) 1974: 20 July (S) , 21 Sept (S,M,P): 1975: 26 Jan (S,M,L,P,A), 26 April (S,M,L,P,A). 48) 1974: 3 Aug (P,A), 21 Sept (S,M); 1975: 5 April (S) Calhoun Co, 49) 1975: 27 March (S,C,A), 6 Sept (S)

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-251Clay Co. 55) 1973: 17 Nov (S.M.L.P); 1974: 4 Jan (S,M,L,P), 23 Feb (S,M,L,P), 4 May (S.M.L.P), 6 June (S,M,L,C), 20 July (S.M.L.P), 31 Aug (S,M,L,P,C), 25 Oct (S,M,L,P), 27 Nov (S,M,L,P,A); 1975: 16 Feb (S.M.L.P.C)," 18 April (S.M.L.P), 21 June (S.L.P.C), 2 Aug (S,M,L,P,C), 19 Oct (S.M.L.P). 56) 1974: 4 Jan (S,M,L,P,C), 23 Feb (S,M,L), 4 May (S,M,L,P,C), 6 June (S,M,L,P,C), 20 July (S,M, L,P), 31 Aug (S,M), 25 Oct (S,M), 27 Nov (S.M.L.P.C); 1975: 16 Feb (S,M,L,P,C,A), 18 April (S,M,L,P,C), 21 June (S,M,P,C), 2 Aug (S,L,P,C), 19 Oct (S,M,L,P,C,A); 1976: 31 Jan (S,M,L,P,C,A) . 57) 1974: 4 Jan (S), 23 Feb (S,L,P), 25 Oct (S) , 27 Nov (S,M,P,A); 1975: 16 Feb (S,M,L,P,C), 18 April (S,M,L,P), 2 Aug (S), 19 Oct (L,P,A); 1976: 31 Jan (S,M,L,P,A). Columbia Co. 58) 1975: 26 April (S) . Duval Co. _70) 1974: 4 Jan (L) , 20 April (S.M.P.C); 1975: 12 March (P.C.A), 4 May (S,M,L,P,C,A) , 1 July (S) , 17 Aug (M.L.P.C.A) ; 1976: 14 Feb (S,M,L,P,C). 11) 1974: 5 Jan (S), 20 April (P) ; 1976: 14 Feb (M) . Escambia Co. 73) 1974: 18 March (S) , 16 June (S,M,P), 7 Aug (S,L,P), 14 Oct (S,L,P,A); 1975: 19 Jan (S,P,C,A), 28 March (S.M.L), 12 June (S,M), 7 Sept (C), 31 Dec (S) . 74) 1974: 16 June (S) , 7 Aug (S) , 14 Oct (S); 1975: 19 Jan (S,L), 28 March (S,M,P,A), 12 June (S), 31 Dec (S); 1976: 18 April (S,M,P). _7_5) 1974: 14 Oct (M) ; 1975: 28 March (S.P.C.A), 12 June (S,L,P), 31 Dec (S,P,C). Gadsden Co. 81) 1973: 18 Dec (S,M,L); 1974: 17 March (S.M.L.P.C), 15 June (S,M,L,P,A); 1975: 27 March (S,M,L,P,C), 10 June (C) , 5 Sept (S). 82) 1974: 6 Aug (M) ; 1975: 18 Jan (S) , 27 March (S.M.L), 10 June (S,M,L), 23 Aug (S,M), 29 Dec (S,M). 83) 1973:

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-25218 Dec (S); 1974: 17 March (S.M.L.P.C); 1975: 18 Jan (S.M.L.P.A), 27 March (S,M,L,P,A), 10 June (S.M.L.P), 24 Aug (S) , 29 Dec (M) . 85) 1975: 30 Dec (S) . 87) 1974: 15 June (M) , 5 Aug (S,L), 12 Oct (S,M,L,P); 1975: 18 Jan (S.M.L.P.A), 27 March (S,M,L,P,C), 10 June (S.M.P.C), 5 Sept (S,M,L), 29 Dec (S.M.L.P.C.A) . 88) 19 75 : 30 Dec (M.L.C); 1976: 17 April (M.L.P.C.A) . 89)1975: 30 Dec (S.M.L.P.C.A). 90) 1974: 17 March (S.M.L), 15 June (S.M.L.P), 6 Aug (S.M.P), 13 Oct (S.M.L); 1975: 18 Jan (S,M), 27 March (S.M, L,P), 10 June (S.M.L.P), 23 Aug (S.L), 29 Dec (S.M.L.P.C.A) . 91) 1975: 18 Jan (S.M), 27 March (S.M.L.P.C.A) , 29 Dec (S) . Hamilton Co. 99) 1974: 8 Aug (M) ; 1975: 1 Feb (S) , 26 April (S,M); 1976: 14 Feb (S). 100) i 97 4: 8 Aug (S.M.L.P.C), 17 Sept (S.M.L, P.C.A), 10 Nov (S,M,L,P); 1975: 23 June (S.M.L.P.C). 24 Oct (S,M, L.P.C.A); 1976: 24 Jan (S.M.L.P). 101) 1975: 1 Feb (S.M.L.P.A), 26 April (S,P). Hardee Co. 102) 1975: 22 March (S.M.L.P.C), 27 May (M.P.C.A), 22 Dec (S,M,L,P,A). 103) 1975: 22 March (S,M,L,P,C,A), 11 Sept (S.M.P.A), 22 Dec (S,M,L,P,C). 104) 1975: 11 Sept (S.M.L.P.C), 21 Dec (S,M, L,P,C,A). Highlands Co. 109) 1974: 24 March (S,M,L,P), 12 May (S.M.L.P.C.A) , ' 23 June (P,A), 29 Nov (S,L,C); 1975: 21 March (S,M,L,P,A), 27 May (S,M,L,P,C), 9 Aug (S,M,L,P,C), 15 Oct (S,L). Holmes Co. 116) 1975: 19 Jan (S.M.L.P.A). 28 March (S,M,P,C), 6 Sept (P). 117) 1975: 28 March (S,P,C), 11 June (S,M,L), 6 Sept (C) , 30 Dec (M). 119) 1975: 19 Jan (S) , 28 March (S), 11 June (S,M 5 L, P,A), 30 Dec (P,C,A); 1976: 18 April (S,M,L,P,C,A) . Jefferson Co. 124) 1974: 16 March (S,M,L,P), 12 Oct (S.M.L.P); 1975:

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-25317 Jan (S.M.L.P.A), 26 March (S,M,L,P,C) , 10 June (S,M,L,P), 23 Aug (S.M.L.P.C), 29 Dec (S) . 125) 1974: 5 Aug (S,M,L,P), 12 Oct (S,M, L,P,A); 1975: 17 Jan (S,M,L,P), 26 March (S,M,L,P,C,A) , 10 June (S,M), 23 Aug (S,M,C), 29 Dec (S,M). Leon Co. 131) 1974: 16 March (S,M,L,P,C); 1975: 26 March (S.M.L.P), 10 June (P,C), 23 Aug (S,M,L,P,C), 29 Dec (S.M.P.C). Liberty Co. 142) 1974: 17 March (S,M,L,P,C,A) ,' 15 June (S,M,), 13 Oct (S,M,L,P,A); 1975: 18 Jan (S,M,L,P,A), 27 March (S,M,L,P,C,A) , 11 June (S,M,L,P,C), 24 Aug (S,M,L,P,C), 30 Dec (S.M.P.C). 144) 1973: 18 Dec (S,M,L,P); 1974: 17 March (S,M,L,P,C,A) , 15 June (S.M.L.P), 6 Aug CS.M.P.C), 13 Oct (S,M,P,A); 1975: 18 Jan (S,M, L,P,C), 27 March (S,M,L,P,C,A) , 11 June (S,P,C), 24 Aug (S.M.L.P, C,A), 30 Dec (S.P.C.A). 146) 1975: 18 Jan (M) , 30 Dec (S,M). 147) 1974: 17 March (S,M,L,P,A); 1975: 18 Jan (S.M.L), 27 March (S,M,L,P,C), 11 June (S,M,L,P,C), 24 Aug (S,M,L,P,C), 30 Dec (S,M, L,C). 148) 1974: 15 June (S,M,L,P), 6 Aug (S.M.L.C), 13 Oct (S,M,L,P,A); 1975: 18 Jan (S.M.L), 27 March (S,M,L,P,C,A) , 11 June (S,M,L,P), 6 Sept (S,M,L,P,A), 30 Dec (S,M,L,P,A). Madison Co. 151 ) 1975: 1 Feb (P) ; 1976: 14 Feb (L) . Manatee Co. 155) 1975: 22 March (S.M.P.A), 27 May (S) , 12 Sept (C) , 22 Dec (S,P). Marion Co. 158 ) 1952: 12 March (S,M W. Beck). Nassau Co. 159) 197d: 4 Jan (S,M,L,P,C), 20 April (S,M,L,P,C), 10 July (S,M), 24 Aug (S,M,L,P), 9 Oct (S,M,P,C), 4 Dec (S,M,L,P,C,A) ; 1975: 12 March (S,M,L,P,C), 4 May (S,M,L,P,C), 1 July (S,P), 26 Sept (S,M); 1976: 14 Feb (S,M,L,P,A). 160) 1974: 20 April (S,M, L,P,C); 1975: 12 March (S,M), 4 May (S,M,P,A), 26 Sept (P) .

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-254Okaloosa Co. 163) 1974: 7 Aug (S) ; 1975: 19 Jan (S) , 28 March (S) , 31 Dec (P). 165) 1974: 18 March (S,M,L,P,C,A) , 16 June (S,M,L,P), 7 Aug (S,M,L,P), 14 Oct (S.M.L.P); 1975: 19 Jan (S) , 28 March (S,M,L,P,C), 12 June (S,M,L,P,C), 31 Dec (S.L.P.C). 166) 1974: 18 March (L,C), 14 Oct (S,M); 1975: 19 Jan (S,M,P,C), 28 March (S.M.C), 12 June (S) . 168) 1973: 13 March (S.P.A K. Tennessen) ; 1975: 29 March (S,M,L), 12 June (S.M.L.P.C), 6 Sept (C) . Orange Co. 169) 1974: 13 July (L.P.C), 28 Nov (S,M,L,P,C); 1975: 15 March (P,C), 31 Oct (S) . JJ4) 1947: 6 and 7 Feb (A H.K. Gouck) . Polk Co. 182) 1975: 22 March (S), 27 May (S,M). Putnam Co. 185) 1974: 14 Feb (P) , 26 March (P) , 23 Nov (P) ; 1975: 15 Jan (M,L). 186) 1974: 19 Jan (L) , 25 May (P) ; 1975: 12 Feb (S,M,P), 18 April (S,P,C). 187) 1974: 19 Jan (S,M,L,P,C), 14 April (S,M,L,P,C,A), 25 May (S,M,L,P,C), 6 July (S,M,L,P), 17 Aug (S,M,L, P,C,A), 6 Oct (S,M,L,P,C,A), 23 Nov (S,M,L,P,C,A) ; 1975: 12 Feb (S,M,L,P,C,A), 18 April (S,M,L,P,C), 26 June (S,M,L,P ,C,A) , 25 Sept CS,M,L,P,A). 189) 1975: 12 Feb (C) , 26 June (P), 31 Oct (S). 191) 1974: 19 Jan (M) , 14 April (M) ; 1975: 5 April (S) . 192) 1974: 25 May (S,M,L,P,C), 6 Oct (S,M,L,P), 7 Dec (S,M,L,P,C,A) ; 1975: 5 April (S,M,L,P,C), 7 Aug (S,M,L,P,C,A) , 31 Oct (S ,M,L,P,C,A) . Santa Rosa Co. 195) 1974: 18 March (S,M,L,P,C), 16 June (S,M,L,P,C), 7 Aug (S.M.L.P), 14 Oct (S,M,L,P,A); 1975: 19 Jan (S,M,L,P,A), 28 March (S.M.L.P.A), 12 June (S,M,L,P,A), 7 Sept (S,M,L,P,C), 31 Dec (S,M,L,P,C); 1976: 18 April (S,M,L,P,C). 196) 1974: 18 March (S,M,L), 16 June (S,M,P), 7 Aug (S,M,L), 14 Oct (S,M,L,P,A); 1975: 19 Jan (S,M,L,P,C,A), 28 March (S,M,L,P,C), 12 June (S,M,L,P,C), 7 Sept (C), 31 Dec (S,F,C).

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-255Seminole Co. ^03) 1974: 21 March (S,M,L,P), 12 May (P) , 28 Nov (P) ; 1975: 15 March (L,P,C), 4 July (S,P). Suwanee Co. 208 ) 1976: 24 Jan (S) . Union Co. 216) 1975: 4 May (A). 217) 1974: 5 Jan (S,M,L,P), 23 Feb (S,M,L,P), 4 May (S,M,L,P,A), 6 July (S,L,P,C), 24 Aug (S,L,C), 9 Oct (C), 6 Nov (S,M); 1975: 12 Jan (S,L,P,A), 8 April (S,M,L, P,C), 1 June (S,C), 5 Oct (S,M,P,C). Walton Co. 220) 1974: 18 March (S,M), 16 June (S,M,P), 7 Aug (S,M,L), 13 Oct (S,M,L,P); 1975: 19 Jan (S,C), 28 March (S,M,L,P,C), 11 June (S,M,L,P,C), 6 Sept (S,L,P,C,A), 31 Dec (S,M,P,C,A). 221) 1975: 28 March (M) . 222) 1974: 6 Aug (P,A). 223) 1975: 19 Jan (S,M, L,P,A), 28 March (S,M,L,P,C,A) , 11 June (P,C,A), 6 Sept (S,P,C,A). Simulium (Simulium) vereoundum Stone and Jamnback Simulium vereoundum Stone and Jamnback, 1955, N.Y. State Mus. Bull. 349: 83 (male, female, larva). Simulium vereoundum, Davies, Peterson, and Wood, 1962, Proc. Entomol. Soc. Ontario 92: 125 (female, male). Simulium vereoundum, Wood, Peterson, Davies, and Gyorkos , 1963, Proc. Entomol. Soc. Ontario 93: 114 (larva). Simulium vereoundum, Stone, 1964, Conn. State Geol. and Natur. Hist. Surv. Bull. 97: 47 (male, female, larva, pupa). Taxonomy. Stone and Jamnback (1955) first separated this species from the closely related S. venustum based primarily on characteristics of the male genitalia. The holotype male was collected in Monroe Co. Pennsylvania and is located in the U.S. National Museum. Stone (1964)

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-256indicates S. verecundum may be the same as S. argyreatum Meigen as defined by Rubtsov but the true identity of S. argyreatum is under question and therefore S. verecundum should be retained as the name for the American specimens. Description . The larva is 6-7 mm long and appears white when recently collected. The head spots are white against a light brown frontoclypeus (Fig. 113). The gular notch is elongate and rounded anteriorly (Fig. 114). The ventral tubercles are visible but not large. The pupa is about 3.5 mm long. The respiratory filaments are 6 in number, almost 3 mm long, and are arranged in pairs. They are spread wide with the bases of filament pairs 3-4 and 5-6 diverging greatly from each other (Fig. 115). The cocoon is brown, slipper-shaped and wellwoven . The male is velvety black in appearance with two silvery, lateral patches on the anterior third of the scutum which connect along the sides of the scutum to a shiny silvery band at the posterior. Each tibia bears a bright white patch. The distimeres are elongate, wide and fairly flat. The arms of the ventral plate in ventral view curve inward slightly toward each other and the median portion or apex of the plate is narrow and elongate in end view (Fig. 116). The wings of the female are 3 mm long. The frons is shiny black. There are ten or more hairs on the underside of the subcosta. The tarsal claw lacks a basal tooth. Each fore tibia has a wide, bright white patch which extends more than three-fourths of the length of the tibia. The ovipositor lobes are dark, sclerotized and distinctly concave on their inner margins, creating an oval space between them (Fig. 117). Distribution . Stone and Snoddy (1959) mention S. verecundum occuring

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-257Figure 113. The head spots of a S. verecundwn larva. Figure 114. The gular notch of a S. verecundwn \t

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-258Figure 115. The pupa and cocoon of S. vevecundvm. Figure 116. Male terminal ia of S. vereaundum.

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-259Figure 117. Terminalia of a female of S. veveoundum.

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-260from Alaska to Nova Scotia south to Washington, Wyoming, South Carolina and Alabama and they add that it is also found in Europe and Northern Asia. Life history. Golini and Davies (1975) report that the eggs of $. vereaundum are .228 mm long and .139 mm wide. Davies et al. (1962) in Canada and Holbrook (1967) in Massachusetts indicate that S. vereaundum eggs overwinter. Larvae appear in late April or early May and adults are present by late May. Anderson and Dicke (1960) found immature stages of S. vereaundum in Wisconsin from July to September. Lewis and Bennett (1973) report larvae develop in 3 weeks and pupae in 4-7 days. Snow et al. (1958) reared one male from a pupa collected in Mississippi on 30 June. Stone and Jamnback (1955) suggest that this species has two or three generations each year and Stone and Snoddy (1969) indicate it is multivoltine in Alabama. Lewis and Bennett (1974) report S. vereaundum first appears in Newfoundland in early June and is present until late October completing three or four generations each summer. Ecology. Davies et al. (1962) indicate that S. vereaundum is generally found in larger streams over 3 m (10 ft) wide. Holbrook (1967) reports that 10 of 14 collections of S. vereaundum were made below pond outlets and swamps. In Massachusetts, streams inhabited by S. vereaundum were very small to about 2.5 meters wide. Snow et al. (1958) collected S. vereaundum from a sandy-bottomed stream. Stone and Snoddy (1969) found this species to be uncommon in Alabama but collected it on the spillways of dams. Lewis and Bennett (1973) found immatures on all available substrates but they were most abundant on trailing vegetation late in the summer. Garris and Noblet (1975) report 23-27%

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-261parasitism by mermithids in S. vereoundum in South Carolina. Dove and McKague (1975) report .001-. 1 ppm of Altosid reduced adult emergence of S. vereoundum by 75-100% at temperatures between 10 and 25°C. Habits . Golini and Davies (1975) report that S. vereoundum females oviposit by first landing at the water's edge on trailing vegetation and then depositing an average of 417 eggs per female in large irregular masses sometimes five layers deep on vegetation. The females of S. vereoundum were found to oviposit more frequently on green and yellow test strips in the flow than on strips of other colors. Stone and Jamnback (1955) and Jamnback (1969) report S. vereoundum does not commonly annoy or attack man. Lewis and Bennett (1973) indicate this species is anautogenous. Abdelnur (1968) captured females feeding on cattle. Florida observations . Stream Parameters Width Depth pH Temperature Velocity Mean: 2.76 m 25.8 cm 5.09 17.4°C (63.3°F) .45 m/sec (1.49 ft/sec) Min: .15 1.27 3.75 6.7 (44) .076 ( .25) Max: 36 166 7.0 27.8 (82) 1.53 (5) Simulium vereoundum is reported for the first time from Florida. Simulium vereoundum was collected from 48 sites in 22 counties (Fig. 118), It was widespread across the northern half of Florida but was not found in collections south of Seminole County. Simulium vereoundum was found in streams from mid-October through mid-June (Fig. 8 ). At Pine Barrens Creek (Site 74) on 14 October larvae and pupae were collected; at all other locations pupae were not found until after October. During June young and mature larvae and pupae were located. There are presently no

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-262-/ .^* Figure 118. Collection locations for S. verecundum in Florida

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-263records of S. verecundum from Florida during July, August, and September. The large whitish and sometimes slightly yellow larvae of this species and the fairly large pupae were most abundant during January through April. Generally, small streams not more than a few meters wide were preferred by S. verecundum, such as Panther Creek, Site 223 (Fig. 119), although a good number of larvae and pupae were found at Pine Barrens Creek (Site 74) which in unflooded condition is about 10 meters across. Other favorable locations including Hatchet Creeic (Site 17), Hogtown Creek (32), Hurricane Creek (83), and a tributary to the Aucilla River (153) illustrate that the suitable depth of the streams varied from about 20 cm to a meter. Optimum stream velocity ranged from .3 m/sec (1 ft/sec) to .76 m/sec (2.5 ft/sec). On one occasion a live pupa from which a female was reared was found in a trickle moving about .076 m/sec (.25 ft/sec). The larvae and pupae were primarily collected in streams acidic in nature but immatures also survived well in flows more neutral in pH reaction. Simulium vereaundum was found on dams or just downstream from impounded water at Shepard's Mill (Site 86), Chattahoochee (88), Tall Timbers (132) and below Lake Melrose (185). Other sites such as Hurricane Creek (83) and Site 90 could be traced to bodies of impounded water upstream. Although a few specimens were found at locations such as Holman Branch (82), Shaw Creek (91) and Camp Branch Creek (118) which were sandy and had little trailing vegetation (even here the larvae were concentrated on the thin sedges or few blades of grass or twigs in the flow), the largest populations were found where eel, other grasses, and additional large and small leafed aquatic vegetation were abundant . Larvae and pupae were found attached to grasses especially and to a wide

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-264Figure 119. Panther Creek, Site 223, where S. verecundum was collected.

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-265range of plants in the streams in addition to twigs and, on a few occasions, to the rock-like clay substrate of a stream. When construction at Sites 6 and 17 destroyed sections of the streams rich in vegetation and forced sampling upstream where the flows continued in a woods where aquatic vegetation was essentially lacking, S. verecundum no longer appeared in the collections. At Kettle Creek (130), the Waccasassa River and Site 196 S. verecundum was never or only rarely discovered in the main flows. Small sandy or sand-mud bottomed drainage ditches which emptied into the larger streams, however, harbored good numbers of S. Verecundum on grass, sedges, dead leaves, shells and ocher substrates. The side flows differed from the main streams in size, velocity, temperature and at Kettle Creek the pH of the side flow was higher than the pH of the main flow. Simuliwn verecundum was associated with 12 other species in Florida's streams (Table 3 ). Most frequently it was found with S. tuberosum and S. slossonae. On 30 occasions it was collected with S. vittatum. Only a few 5. verecundum larvae with signs of protozoan infections have been collected and no mormithids have been noted in the imma tures . A character given in some taxonomic keys to separate S. verecundum larvae from its relative S. venustum is that S. vcrecu.nd.urn possesses more than 52 rays in each cephalic fan. Few of the S. verecundum larvae collected in Florida have more than 52 rays and most range from 45-50. Also an unexpected fulvous area lines the gular notch on some specimens and others have an apparently uncharacteristic reddish abdominal tinge. The adults reared or dissected from immatures in populations with variant characters such as these have all proven to be S. verecundum.

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-266Florida collection records for S. verecundum. Alachua Co. 1) 1973: 7 Dec (S) , 13 Dec (P) ; 1974: 2 Feb (M,L,P), 12 April (S,M), 5 May (P) , 30 Oct (S,L), 23 Nov (M,L,P,A); 1975: 10 Jan (S,M), 12 Feb (S,L,P). 6) 1973: 17 Nov (S) , 7 Dec (S,M); 1974: 2 Jan (M,P,A), 2 Feb (S.M.P), 7 March (S). 15) 1975: 31 Jan (P). 17) 1974: 2 Feb (S,M), 7 March (M.L.P), 12 April (S.M.L), 5 May (S.M.L.P), 6 June (S,M). 18) 1974: 25 Oct (M) ; 1975: 10 Jan (S), 31 Jan (S,L,P), 8 April (S,M,L), 11 May (S) , 14 June (S) . 32) 1974: 2 Feb (S,M,L,P,A), 9 March (S,P), 13 April (M,P,C,A); 1975: 10 Jan (S,M,L,P,A), 4 April (S,M,L,P,C). 39) 1974: 25 Oct (M). Baker Co. 41) 1974: 4 Jan (S) , 6 Nov (S). Bradford Co. 42) 1975: 19 Nov (S) . 47) 1975: 26 Jan (S,M,L), 26 April (S.M.P). 48) 1975: 26 Jan (P,A). Dixie Co. 68) 1975: 17 Jan (S) , 26 March (S,M,P,C,A). Duval Co. 72) 1974: 20 April (S) ; 1975: 12 March (S,M,L,P,A), 4 May (S); 1976: 14 Feb (S) . Escambia Co. 73) 1974: 16 June (M) ; 1975: 12 June (S) . U) 1974: 16 June (S), 14 Oct (S.M.L.P) ; 1975: 19 Jan (S.L.P.A). Gadsden Co. 81) 1973: 18 Dec (M) ; 1974: 17 March (S,M); 1975: 27 March (S.M.P.A). 82) 1975: 18 Jan (L) . 83) 1974: 13 Oct (S) ; 1975: 18 Jan (S,M,L,P), 27 March (S,M,L). 86) 1975: 18 Jan (P,C, A). 87) 1974: 15 June (M.L.P); 1975: 18 Jan (S,M,L,P), 27 March (S,M,P), 29 Dec (S.L.P.A). 88) 1975: 30 Dec (P,A). 90) 1974: 17 March (S,M,L,P); 1975: 18 Jan (S,M,L,P,C,A) , 27 March (S,M,L,P, C,A), 10 June (S) , 29 Dec (S,M). 91) 1975: 27 March (P,A).

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-267Hamilton Co. 101) 1975: 1 Feb (S,M,L,P,A), 26 April (S). Holmes Co. 118 ) 1975: 18 Jan (S,M). 119 ) 1975: 19 Jan (S,M). Jefferson Co. 124) 1975: 26 March (S) . Lafayette Co. 130 ) 1975: 17 Jan (S,M,L,P,C,A) , 26 March (S,M,L,P), 28 Dec (S). Leon Co. 131 ) 1974: 16 March (S,M,L); 1975: 26 March (S,M,L,P,C), 29 Dec (S). 132) 1974: 16 March (S,M,P); 1975: 17 Jan (S) . Levy Co. 138 ) 1975: 23 March (P). 139) 1975: 23 March (S,M,P,C,A). 141) 1975: 23 March (M) . Liberty Co. 142) 1975: 18 Jan (S,M), 27 March (S,M). 147 ) 1974: 17 March (S,M,P,A); 1975: 18 Jan (S) , 27 March (S,M). Madison Co. 153 ) 1974: 16 March (S.M.P.C); 1975: 17 Jan (S.M.L.P), 26 March (S,M,L,P,C,A) , 10 June (P) , 29 Dec (S,M,P,C,A). Nassau Co. 160 ) 1975: 4 May (S) . 161 ) 1975: 12 March (S,P). 162 ) 1975: 12 March (S,P). Putnam Co. 185 ) 1974: 26 March (S,M), 18 May (S,M); 1975: 15 Jan (S.M.P), 4 April (S,M,L,P,C), 11 May (S,M,L,P). 186 ) 1974: 19 Jan (S), 14 April (M,L,P,A), 25 May (S,M,P), 23 Nov (M,P,C); 1975: 12 Feb (S,M,L,P,C,A), 18 April (S,M,L,P,C). Santa Rosa Co. 196 ) 1974: 18 March (S,M,L,P,A), 14 Oct (S,M); 1975: 28 March (S,M). Seminole Co. 203 ) 1975: 27 April (M) . Taylor Co. 210 ) 1973: 17 Dec (S,M,L,P,C); 1974: 16 March (S,M,L,P); 1975: 17 Jan (S), 28 Dec (S,M,P,C). 213 ) 1975: 17 Jan (S.M.L, P,C), 26 March (S,M,P,C,A). 215 ) 1975: 26 March (S,M,P,C,A). Union Co. 217 ) 1974: 5 Jan (S) ; 1975: 12 Jan (L.P.A), 8 April (S). Walton Co. 223 ) 1975: 19 Jan (S.L.P), 28 March (M) .

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-268Cnephia r.pccies Undetermined No. 1 On 13 March 1969 in a small stream which passed under the Millhopper Rd. (State Road 232) .8 km (.5 mi) southeast of the Devil's Millhopper, Gainesville, Alachua County (Fig. 122), Mr. L. Goldman collected larvae of a black fly species which has been found nowhere else in Florida. The larvae with dark histoblasts are 6-7 mm long and appear pinkish brown in alcohol. The head spots are rather indistinct with only the two elongate, dark brown medial groups of spots very visible (Fig. 120). The antennae have dark brown basal segments, lighter brown second segments, and are very thin the rest of their lengths which extend beyond the cephalic fan stalk. The cephalic fans contain a large number of rays, about 80-90. The gular notch is a very tiny inverted-V (Fig. 121). The submenturc has large broad lateral teeth and a shorter median tooth. The respiratory histoblast contains 20 to 30 long thin filaments which rise first in groups of 4 and 6 and then bifurcate in pairs off thicker, elongate stems. The abdomen in dorsal view bulges at the fifth segment, is widest at about the seventh segment, and tapers thereafter. There are about 64 rows of anal hooks with 10-12 hooks-row. The anal sclerite is elongate or tall with very short, dark, thin anterior and posterior arms. The anal tubercles are large. The. subinentum of this species approaches that of Cricpliia mutata (Malloch) and the size and shape of the throat cleft are somewhat similar to that of C. mutata but other characters disagree strongly such as the number of respiratory histoblast filaments and the shape of the anal sclerite. The Goldman collection site in recent years has been disrupted by a housing development and no stream was observed to flow at the collection localit) during this research.

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-269* • • i • Figure 120. Cephalic apotome of a larva of Cnephia species No. 1. Figure 121. Venter of the head capsule of a larva of Cnephia epeoies No. 1.

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-270Figure 122. Collection location for Cnevhia species No. 1 in Florida.

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-271Simuliion species Undetermined No. 1 On 19 January 1975 in the vicinity of Pine Barrens Creek (Site 74), Escambia County (Fig. 126), a pupal exuvium with cocoon was collected which is unlike any other black fly specimen found in Florida. The specimen, shown in Fig. 123, bears respiratory organs consisting of four filaments each rising in two pairs off short petioles. The long, thin filaments are about 3 mm in length. The cocoon is light brown, wellwoven, has anterior edges which are concave in lateral view and bears a short mid-dorsal anterior projection. The cocoon is 3 mm long on the dorsum and 3.5 mm long ventrally. The specimen was collected from pebbles or dead leaves in a small 25 cm wide, 2 cm deep side flow to Pine Barrens Creek (Fig. 127). The small stream emerged from a woods on a sand and round-pebble bed and flowed at .61 m/sec (2 ft/sec). The water temperature was 17.5°C (63.5°F) and the pH was 4.5. Simulium tuberosum larvae were also present in the small stream. A medium-aged larva with incompletely-developed histoblasts was collected under the same conditions as the exuvium and is believed to represent the same species. The larva is 5 mm long and has a light brown head capsule and abdomen. The head spots appear dark brown on a lighter yellow brown cephalic apotome (Fig. 124). The posterior medial group of spots expands posteriorly. The antennae extend just beyond the stalk of the cephalic fan. The cephalic fans each consist of about 60 rays. The gular notch is longer than wide, rounded anteriorly and extends slightly less than halfway to the teeth of the submentum (Fig. 125) The submentum has prominent lateral teeth which are nearly as large as the conspicuous medial tooth. The anal sclerite has long anterior arms.

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-272The anal tubercles are prominent. The specimens appear to represent the subgenus Eusimulium and are similar but not identical to S. alarkei Stone and Snoddy, differing in the length of the filament petioles, and S. latipes (Meigen) , which is a more northern species and has a long antero-dorsal projection on the cocoon. Attempts to collect additional specimens of this species during December 1975 and April 1976 were futile.

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-273Figure 123. Pupal exuvium and cocoon of Simuliwn species No. 1,

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-274Figure 124. Larval head spots of Simulium species No. 1. Figure 125. Venter of the head capsule of the larva of Simulium i No. 1. pec

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-275rJ J&* Figure 126. Collection location for Simuliwn species No. 1 in Florida,

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-276Figure 127. Small flow to Pine Barrens Creek, Site 74, where Simulium species No. 1 was collected.

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Leuoocytozoon smithi Transmission Table 5 presents the results from the sentinel turkey exposures. On four occasions one bird of each triplet exposed in Alachua County during May-July 1975 became infected with L. smithi. The presence of the disease and vectors indicated by the positive sentinels at the Hatchet Creek Preserve during May and July was further supported by blood smears positive for L. smithi which were obtained from two semiwild turkeys (1/4 wild, 3/4 domestic) being reared in an outdoor pen at the Preserve. The first few positive sentinels (9170, 223 and 245) served as initial donors for lab transmission experiments. All subsequent lab transmission donors were lab-infected birds except during early 1976 when two sentinels, 483 and 485, were used as donors. Four turkeys infected during the summer and fall of 1975 were held over the winter to serve as donors for transmission trials early in 1976 if difficulties were encountered in 1976 as they had been in 1975 in acquiring younger, more manageable birds to field infect during January through March. It was hoped that an increase of the Leuoocytozoon smithi infection reported in some Leuoocytozoon species (Desser et al.,1967; Cook, 1971; Fallis et al., 1974) when the mating season began or under conditions of stress would occur in the overwintering laboratory birds. Mating usually begins during January or February with turkeys, and eggs were found frequently in the holding room beginning in February. Two of the female -277-

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-278Table 5 . Sentinel turkey locations and results. Turkey Exposure Number Unit Location Dates Leucocytozoon 142, 143, cage Fisheating Creek (Site 95) 21-22 April 1975; 30-31 May 1975 negative 149, 150, ramp trap 175 Lochloosa Creek (Site 22) 29-30 April 1975 negative 9168, 9169, ramp trap 9172 SR 225/340 Junction (Site 8) 9-11 May 1975 negative 9163, 9167, cage 9170 Hatchet Creek Preserve (Site 24) 17-23 May 1975 only 9170 posit ive , 2 June 245, 246, cage 247 SR 225/340 Junction 16-22 June 1975 only 245 positive, 11 July 223, 224, cage 312 Lochloosa Creek 2-10 July 1975 only 223 positive, 20 July 313, 314, ramp trap 315 Hatchet Creek Preserve 3-10 July 1975 only 315 positive, 20 July 486, 487, cage 488 Hatchet Creek Preserve 18-25 Feb 1976 negative 479, 480, cage 481 Lochloosa Creek 4-8 March 1976 (birds stolen) 476, 477, cage 478 SR 225/340 Junction 4-8 March 1976 (birds stolen) 483, 484, cage 485 SR 225/340 Junction 9-16 March 1976 483, 484, 485 all positive, 22 March

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-279birds (223 and 325) were severely wounded on the posterior lateral regions of their bodies during the mating attempts of a huge torn turkey initially housed with the females. Despite the flow of sex hormones and the stress of egg laying, mating and mating wounds, and very cold nights spent with little significant artificial heat, as of early March no significant rise in the gametocytemias of the birds was observed and they were sacrificed. One bird (324) previously exposed in the field on 7 January and fed upon by several black flies had a noticeable but short-lived rise in its gametocyte count approximately two weeks after the field exposure. Temperatures during the dav from 10-i5°C (50°F) and at night around 0°C (28-35°F) which required that the sentinel poults be brought in at night might explain the negative results in the 18-25 February 1976 exposure. Although turkeys and equipment were stolen from the sentinel sites during the important first week in March 1976 when spring species such as S. congavaenavvjn and C. oraithophitia were flying, the three heavy infections achieved in sentinels the following week compensated for the losses. One of these sentinels (483) with a heavy gametocyte level of 1-2 gametocytes per oil immersion field was the only turkey observed to die from other than preplanned causes. It cannot be stated for certain that L. strn'thi was the reason for its demise. Positive infections were obtained in this research in sentinels exposed primarily on the ground, on a high stream bank (Fig. 2 ) or on a platform 1.53 m above the ground. Observation of the efforts of Dr. D.J. Forrester in 1974 at Fisheating Creek revealed that sentinel birds placed in cages in cypress trees 6 m (20 ft.) above the ground and exposed for two week periods during the summer became positive for L. cmithi but poults

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-280placed in similar cages at ground level remained negative for the disResults from the Manitoba trapping were presented in Table 4 and the results from the ramp, blackout box, and exposed turkey collecting are presented in Tables 6, 7 and 8. The Manitoba captures revealed that a number of species were on the wing including species in the Pkoaterodovos group, for example at Lochloosa Creek where immature populations of a number of the Phosterodoros species occurred. The ramp trap and blackout box collections revealed the presence of only a single species, S. slossonae. The blackout traps were used on clays when temperatures ranged from 25-22. 2°C (77-90°F) and relative humidities ranged from 5890% with the higher humidities most often encountered. Each successful trapping period yielded 1 to 16 specimens of S. slossor.ae and when an infected host was used as the attractant, blood-fed flies which were collected provided a starting point for early lab transmission trials. While the birds were uncovered in the blackout box trials it was noted that they were extremely sensitive to the presence of even a single black fly. They reacted by shaking, rubbing or pecking, which prevented many flies from feeding and killed numerous flies. These difficulties might have been overcome by restraining the host in some manner, but it was found that even the most docile restrained birds became very agitated and, if on their sides, struggled vigorously to stand to move their bowels or for some, other reason. It was also observed that: a number of black flies landed and fed on the head, neck or legs of the host. Some crawled beneath the feathers and emerged a few minutes .later biood red and quickly flew off while the turkey was still in the uncovered condition. The ramp trap was hoped to be an improvement

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•281Table 6 . Black flies captured in ramp traps. Turkey Number Location Dates Captures (all 1975) (all females) 9170 Rt 225/340 June21-22 June tion 149, 150, Lochloosa Creek 29-30 April 175 (Site 22) 9172, 9168, Rt 225/340 June9-11 May 9169 tion (Site 8) 9170 ' Hatchet Creek 18-20 June Preserve (Site 24) 1 S. slosi 313, 314, Hatchet Creek 3-10 July 315 Preserve 2 S. slossonae

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-282Table 7 . Blackout box trapping results, Turkey Number Location 121 121 121 121 121 121 121 121 121 9170 9170 9170 223, 224, 225 9170 9967, 9947 Locbloosa Creek (Site 22) Lochloosa Creek Lochloosa Creek Lochloosa Creek Lochloosa Creek Tributary to Lochloosa Creek (Site 23) Stream from Lake Melrose (Site 185) SR 225/340 Junction (Site 8) Sandy Hatchet Creek (Site 1) Flow to Newnan's Lake (Site 18) Hatchet Creek (Site 20) Turkey Creek (Site 216) Lochloosa Creek Double Run Creek (Site 43) Fisheating Creek (Site 95)" n . -,.. Captures Date lime * (all females) 15 Aug 1974 1730-1845 S. slossonae 20 Aug 1755-1930 S. slossonae 27 Aug 1800-1930 S. slossonae 28 Aug 0730-0930 S. slossonae 12 Sept 1620-1945 S. slossonae 25 Sept 1550-1900 S. slossonae 14 May 1975 1430-1830 3 June 0900-3130 S. slossonae 3 June 1140-1230 S. slossonae 3 June 0830-0930 3 June 1000-1215 15 June 1740-2020 S. slossonae 2 July 1700-1730 6 July 0830-0940 S. slossonae 18 Sept 0900-1.130

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-283Table 8. Black fly captures from exposed turkeys. Turkey Number Exposure Location Date Time Captures 142, 143 Fisheating Creek (Site 95) 142, 143 Fisheating Creek 313, 314, 315 9170 9170, 245 223 9947, 9967 9969 316, 317 Hatchet Creek Preserve (Site 24) Double Run Creek (Site 43) Sandy Hatchet Creek (Site 1) Sandy Hatchet Creek Fisheating Creek Lochloosa Creek (Site 22) 320,

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-284in that it still used turkeys as attractants; however, flies entering to feed were considered captured and the col lector could recover specimens as his schedule allowed. The ramp trap used in Alachua Co. and the 4 to 5 ramp traps used in Glades Co., primarily in another study for five 2-week periods during May-October 1975, captured many mosquitoes and other insects but only three black flies were recovered in Alachua Co. and none in Glades Co. The black flies captured, being the ornithophilic species S. slossonae, were significant and gained in importance when one of the sentinel birds (315) used during the 3-10 July 1975 exposure period when two S. slossonae were recovered from the trap developed leucocytozoonosis. However the success of capturing simuliids with ramp traps overall was poor. The possible inappropriateness of the ramp trap for capturing black flies was suggested during a little experiment in June 1975 that was conducted behind the Veterinary Entomology lab. At the slanted entrance slit of the ramp trap four reared S. jonesi females were released. Three of the four flies released flew to the upper screened portion of the trap opposite from the entrance slit where the release occurred. One black fly flew a short loop out of the vial into the trap and landed halfway down the slanted ingress screen, clinging upside down. It immediately commenced the typical black fly walking and wandering behavior observed often in the collecting jars of the Manitoba traps. The black fly walked up the slanted entrance ramp on the inside to the metal frame securing the organza and immediately flew out the narrow 2.5 cm (1 in) entrance slit: and escaped. Downe and Morrison (1957) reported that S. vittatm and G. desorum readily entered animal baited traps through baffle entries but mentioned, once the flies were inside, they ignored the host and searched for a means to escape.

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-285Perhaps this behavior plus reduced visibility of target animals or restricted flow of host attractant odors through the wood and fine-meshed polyester organza detracted from the effectiveness of the trap. The flies captured from the exposed turkeys reveal the presence during January and March of a second ornithophilic species, S. aongaveenavum, which was observed to feed on the turkeys. Black flies approached the turkeys on days when the temperatures ranged from 22.2°C (72°F) to 30.6°C (87°F) and the wind was calm or blew in gusts up to 16 km/hr. Rarely were situations encountered where more than two or three flies of any species were attracted to or were observed feeding on one turkey at the same time. Circumstances were never experienced such as Garris et al. (1975) report where several hundred black flies were observed around a single turkey in one half hour period. It was noted that, while black flies were in flight and were collected from exposed turkeys during almost all periods during the day, the number of simuliids around exposed turkeys and around my head increased with the approach of thunderstorms during the summer afternoons. Black flies were observed to bite and feed on turkeys in the field and in the lab at a number of body regions. Biting was confirmed when a stationary fly in the head-down feeding position was observed to pull its mouthparts out of the skin and blood feeding was noted by a hematoma on the skin and bright red blood in the gut of the dissected fly. In the field flies were observed to crawl beneath the feathers of the wing and the lower abdominal region to feed and also fed on the top of the head, on the neck, and on bare portions of the legs. In the lab positive feedings were achieved by placing flies against the breast and bare underside of the wing as well as on the neck and head. Compared to the

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-286numbers of flies of other species thaL fed on turkeys in the field or in the lab many more S. slossonae were observed to feed on the birds. Both wild-caught and reared specimens of S. slossonae and S. aongcveewmm fed on turkeys. Reared S. congareenarum females were very reluctant to feed on the turkeys, regardless of whether they were young or old fixe,, were provided with a sugar source or deprived of food for 24-48 hours, were provided with water or deprived of water for 3-12 hours, held with males or held alone or even taken into the field in hopes of stimulating feeding. Reared, non-blood fed S. congareenaruin dissected during January 1976 were discovered to have large, well-developed eggs which suggests at least some autogenous egg development. Six reared C. ornithophilia fed on the turkeys as did six wild-caught S. neridicnale.. No S. mevidionale adults were reared. In addition, reared females of the S. (Phostevodovos) species jenningsi, jonesi, and notiale and undetermined wildcaught S. (Phosterodoros) species fed on the turkeys. Simdium tuberosum would not feed on turkeys in the lab. In the lab flies fed on turkeys during all hours of the day, especially during the late afternoon, and even when it was dark outside. Black £ U e S stimulated to feed usually went right to the turkey when the feeding vial was placed against the bird and were biting within five minutes. Only rarely was feeding stimulated when flies reluctant to feed were kept on the turkeys thirty minutes or longer. Garris et al. (1975) observed black flies feed to such repletion on turkeys that they could not fly and would fall to the ground. During two years of feeding black flies on turkeys under many different circumstances I have noted that once the black flies become replete with blood they remove their mouth parts and always quickly fly away if unrestrained. If confined in a feeding vial

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-287they exhibit for a short period a burst of flight activity after which they settle down and become quiescent for a number of hours. Simuliwn slossonae was observed to feed under a variety of temperatures (13 -32°C 55 90°F) and relative humitidies (40-90%). Uninterrupted feeding times for S. slossonae ranged from 1 to 10 minutes with an average of 4.3 minutes. Simuliwn congareenar-um fed at temperatures from 17-27°C (62-80. 5°F) and at relative humitidies of 60-87%. Simulium congar eenarum feeding times ranged from 1.5 to 5 minutes with most being 3-5 minutes. On a number of occasions in the field and lab a small droplet of blood was observed to protrude from the anus of S. slossonae just prior to terminating feeding. Most black flies fed to repletion rapidly, of ten in three to four minutes, and were not readily disturbed while engorging. Complete engorgement occurred most swiftly, in as little time as one minute, when flies fed on the posterior portion of the head or on the neck. Longer feeding periods were apparently spent attempting to create a pool of blood beneath the skin. Actual engorgement marked by a swift ballooning of the fly's abdomen as the gut filled with blood occurred rapidly, in about one minute. One S. congaveenavwn aspirated out of a Manitoba trap in the field was placed against a turkey's neck while still in the aspirator tube. The fly walked to the small neck feathers and clinging to the top side of a feather with only the proboscis touching the skin of the turkey proceeded to engorge in three minutes (1729-1732 hrs) . Cnephia ovnithophilia, a large fly, took a larger blood meal and required a longer feeding period, average 8.5 minutes, than usual for the other species observed. One Cnephia female propped itself up in the typical feeding position for this species which is at a 75 to 80° angle almost

PAGE 301

-288perpendicular to the host's body surface, inserted the mouth parts at 1550 hrs and remained attached, not filling with blood and releasing until 1606 hrs. The hematomas created by feeding Cnephia on the chest of a turkey, often 3 mm in diameter, were usually larger than the feeding marks caused by other species. On 25 March a 5 mm hematoma resulting from a 4 minute feeding by S. congareenavvm was noted. Simulim meridionale fed within 2 to 8 minutes and seemed to engorge more extensively than any other species with the abdomen expanding to about three ti raes the size of the thorax. Table 9 presents a summary of the successful transmission trials. Three black flies.S. slossonae, S. oongareenarm and S. meridionale. all proven as vectors in other States, were found to be pathophorous agents of L. smithi in turkeys in Florida. Of 116 S. slossonae that fed once on turkeys, 26 fed at least twice once on an infected bird and one or more times on clean turkeys and 15 positive transmissions occurred. Most transmissions were achieved by the bite of a single fly. Two wild caught S. congaveenarwi fed directly on a clean turkey with no leucocytozoonosis resulting. Of 7 S. congareenarum that fed once on infected birds, 3 of the 4 that survived fed a second time, on clean birds, and 3 transmissions occurred. With S. meridionale 6 females fed one time on infected turkeys, 1 fly fed for a second time and £< ^.^ ^^ .^ was achieved. Injections of possibly infected flies ground and innoculated in physiological saline intraperitoneal^ into turkeys were tried on four occasions with S. slossonae and on one occasion with S. meridionale with negative results. Two females of S. noiiale and one of S. taxodium fed once on infected hosts but died soon thereafter or would not feed again on uninfected hosts. One S. jenningsi female, one

PAGE 302

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-290S. jonesi, and three C. omithophilia fed twice but no L, smithi infections resulted. The vectors listed in Table 9 were collected from infected turkeys exposed in the field (field exposure), captured in Manitoba traps, were reared, or were netted. The collection locations of the vectors include 10 different sites in a total of 7 west, north, central and south Florida counties. The black flies were experimentally infected during nearly every month of the year with most transmissions occurring during late summer-early Fall (August-early October) and in the Spring (MarchApril). Flies held for 2-7 days after taking an infected blood meal transmitted the disease. Transmissions were achieved with feeding times that ranged from about 1 minute in interrupted feedings to 10 minutes for some full engorgements. One reared 5. slossonae infected on 30 September transmitted L. smithi to host #340 on 2 October in an interrupted feeding and the same fly transmitted L. smithi to turkey #342 the next day, 3 October, during a more complete feeding. Recipients for all transmissions were young turkeys from 18 days to 19 weeks old, with most being 3-7 weeks of age. The prepatent period from the transmission date when an infected fly bit the clean turkey to the appearance of gametocytes (Fig. 128) in the blood smears was 10-15 days. Deeply staining ookinetes of L. smithi (Fig. 129) 20-22 y long and 2.5-3.5 v wide with large crystalloid vacuoles have been noted in smears of the gut contents of S. slossonae 11 hours after a blood meal was taken from an infected turkey and a little later in the gut contents of infected S. congareenarwn. Spherical hyaline structures on the midgut believed to be oocysts were observed many times in flies that had fed on infected turkeys but unruptured structures displaying distinct

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-291Figure 128. Gametocytes (G) of L. smithi among normal turkey blood cells. Figure 129. Ookinetes of L. smithi.

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-29220u Figure 130. Sporozoites of L. smithi photographed in sal: figure 131. Stained sporozoites of L. smithi.

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-293elongate sporozoites packed like toothpicks as illustrated for other species of Leucooytozoon (Pallia and Bennett, 1961a) were never positively located. Sporozoites (Fig. 130 and 131) about 10-15 v long and 1.25-2 V wide were observed issuing in a saline preparation from the gut region, probably from squashed oocysts, 45 hours after the female S. slossonae fed on a heavily infected turkey. In a S. slossonas female that had taken an infected blood meal 72 hours prior to dissection 50-100 sporozoites were observed escaping into the saline from the salivary glands and only a few sporozoites were noted in the gut region. The sporozoite stage with a blunt or rounded and wider anterior end and a tapering posterior end appeared slightly longer and wider in a fresh saline preparation in contrast to the fixed and stained preparations. Sporozoites located on the slides of guts and salivary glands of flies which transmitted L. smithi to turkeys were normally few in number except in one instance when the vector was allowed to insert its proboscis for only 45 seconds. The salivary glands of this fly were found to contain many sporozoites. From the above investigations it is concluded that Simulium slossonae, the most widely distributed species of the three vectors and the only species present year round is the most important vector of L. smithi to turkeys in Florida. Simulium aongaveenavum is a secondary vector of significance especially during the first three or four months of each year. In the counties bordering the Apalachicola River, during the two to three week mass emergence' period of S. meridionale in April and during most of May when the adults are still flying, S. meridionale may be an important vector of L. smithi.

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CHAPTER V CONCLUSIONS 1) From this research it is concluded that sixteen previously described species representing two genera and six subgenera of black flies occur in Florida. In addition, two species not positively identified and possibly previously undescribed are recorded from the State. Eight of the known species and the two undetermined species are reported for the first time from Florida. 2) Species such as S. slossonae and S. tuberosum are widely distributed throughout much of the State, are adapted to a wide range of lotic habitats and are present all year. Certain species such as S. dixiense, S. haysi, and S. notiale have a limited distribution in Florida and occur at only a few locations. Other species such as C. ovnithopkilia , S. congareenarum , and 5. vereoundion were found to have a more limited seasonal occurrence and were undetected during the warm summer months. 3) Except during mass emergences of S. mevidionale in April along the Apalachicola River, black flies rarely cause harm directly to man in Florida. The potential for the transmission of diseases, especially viral encephalitis, by simuliids nevertheless exists. 4) Three species of black flies already recognised as vectors in other areas of the U.S. were incriminated through lab and field efforts 294-

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-295as vectors of L. smithi to turkeys in Florida. In order of decreasing importance, these species are: S. slossonae , S. oongareenarum, and S. meridionale. 5) Barring major breakthroughs in prophylatlc medications or vector control, the widespread distribution of the ornithophilic vectors of L. smithi would restrict any potential turkey industry to locations miles away from black fly streams or force screened outdoor or indoor rearing procedures.

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LITERATURE CITED Abdelnur, O.M. 1968. The biology of some black flies (Diptera: Simuliidae) of Alberta. Quaest. Entomol. 4(3): 113-174. Agricultural Situation. November 1974. Tomorrow's turkeys. 58(10): 2-4. Aikawa, M. , and C.R. Sterling. 1974. Intracellular parasitic protozoa. Academic Press, New York. 76 pp. Anderson, J.R., and G.R. DeFoliart. 1961. Feeding behavior and host preferences of some black flies (Diptera: Simuliidae) in Wisconsin. Ann. Entomol. Soc. Amer. 54: 716-729. Anderson, J.R., and G.R. DeFoliart. 1962. Nematode parasitism of black fly (Diptera: Simuliidae) larvae in Wisconsin. Ann. Entomol. Soc. Amer. 55: 542-546. Anderson, J.R., and R.J. Dicke. 1960. Ecology of the immature stages of some Wisconsin black flies (Simuliidae: Diptera). Ann. Entomol. Soc. Amer. 53: 386-404. Anderson, J.R., V.H. Lee, S. Vadlamundi, R.P. Hanson and G.R. DeFoliart. 1961. Isolation of eastern encephalitis virus from Diptera in Wisconsin. Mosquito News 21(3): 244-248. Anderson, J.R., and J. A. Shemanchuk. 1975. Maintenance and transportation of black fly (Diptera: Simuliidae) larvae in nenagitated water. Ann. Conf. Calif. Mosquito Control Ass. Proc. Pap. 43: 120-122. Anderson, R. I . , L.E. Fazen, and A. A. Buck. 1975a. Onchocerciasis in Guatemala. II. Microfilariae in urine, blood, and sputum after dicthylcarbamazine. Amer. J. Trop. Med. Hyg. 24(1): 58-61. Anderson, R.I., L.E. Fazen, and A. A. Buck. 1975b. Onchocerciasis in Guatemala. III. Daytime periodicity of microfilariae in skin. Amer. J. Trop. Med. Hyg. 24(1): 62-65. Anthony, D.W. , and D.J. Richey. 1958. Influence of black fly control on the incidence of Leucooytozoon disease in South Carolina turkeys. J. Econ. Entomol. 51(6): 845-847. Atchley, F.O. 1951. Leucom/tczoon andrewsi n. sp. from chickens observed in a survey of blood parasites in domestic animals in South Carolina. J. Parasitol. 37: 48.3-488. 296-

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-297Bailey, C.H. , R. Gordon, and J. Mokry. 1974. Procedure for mass collection of mermithid postparasites (N'ematoda: Mermithidae) from larval blackflies (Diptera: Simuliidae) . Can. J. Zool. 52: 660-661. Barreto, P. 1969. The species of black flies found in Colombia (Diptera; Simuliidae). N.Y. Entomol. Soc. J. 77(1): 31-35. Batson, B.S., M.R.L. Johnston, M.K. Arnold, and D.C. Kelley. 1976. An iridescent virus from Sinrulium sp. (Diptera: Simuliidae) in Wales. J. Invert. Pathol. 27(1): 133-135. Beck, W.M. , Jr. 1965. The streams of Florida. Bull. Fla. State Mus. Biol. Sci. 10(3): 126 pp. Bennett, G.F. 1960. On some ornithophilic bloodsucking Diptera in Algonquin Park, Ontario, Canada. Can. J. Zool. 38: 377-389. 32 Bennett, G.F. 1963a. Use of P in the study of a population of Simuliwn rugglesi (Diptera: Simuliidae) in Algonquin Park, Ontario. Can. J. Zool. 41: 831-840. Bennett, G.F. 1963b. The salivary gland as an aid in the identification of some simuliids. Can. J. Zool. 41: 947-95?. Bennett, G.F., A.G. Campbell, and M. Cameron. 1974. Hematozoa of passeriform birds from insular Newfoundland. Can. J. Zool. 52(6): 765-772. Bennett, G.F., and A.M. Fallis. 1971. Flight range, longevity, and , habitat preference of female Simulivm eupyacbuinieulum Davies (Diptera: Simuliidae). Can. J. Zool. 491,9): 1203-1207. Bennett, G.F., P.C.C. Garnham, and A.M. Fallis. 1965. On the status of the genera leucocytozoon Ziemann, 1898, and Haonopi'oteus Kruse, 1890 (Haemosporidia, Leucocytozoidae and Haetnoproteidae) . Can. J. Zool. 43: 927-932. Bennett, G.F., M. Laird, R.A. Khan, and CM. Herman. 1975. Remarks on the status of the genus Leucoaytozoon Sanihon, 1903. J. Protozool. 22(1): 24-30. Berner, L. 1950. The mayflies of Florida. Univ. Fla. Biol. Sci. Ser, 4(4): 267 pp. Bierer, B.W. 1954. Buffalo gnats and Leucoaytozoon infections of poultry. Vet. Med. 49: 107-110 and 115. Blickenstaf f , C.C. 1970. Common names of Insects. Committee on Common Names of Insects, Entomological Society of America. 36 pp. Borg, K. 1953. On Leucocytozoon In Swedish Capercaillic, Black Grouse and Hazel Grouse. Berlingska Boktryckeriet, Lund: 1-109.

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-298Bradbury, W.C., and G.F. Bennett. 1974a. Behavior of adult Simuliidae (Diptera). I. Response to color and shape. Can. J. Zool 52251-259. Bradbury, W.C. , and G.F. Bennett. 1974b. Behavior of adult Simuliidae (Diptera). II. Vision and olfaction in near-orientation and landing. Can. J. Zool. 52: 1355-1364. Brues, C.T., A.L. Melander, and F.M. Carpenter. 1954. Classification of insects. Bull. Mus. Comp . Zool. 108: 917 pp. Burton, G.J. 1971. Attachment of first instar Sinralium darrrnoswn (Diptera: Simuliidae) larvae to older larvae. Mosquito News 31(3): 443. Byers, C.F. 1930. A contribution to the knowledge of Florida Odonata. Univ. Fla. Biol. Sci. Ser. 1(1): 327 pp. Byrd, M.A. 1959. Observations on Leuaooytozoon in pen-raised and free-ranging wild turkeys. J. Wildlife Management 23: 145-156. Cameron, A.E. 1922. The morphology and biology of a Canadian cattleinfesting black fly, Sirrraliwn simile Mai. (Diptera, Simuliidae). Dora. Can. Dep. Agr. Bull. 5: 1-26. Carestia, R.R., R.L. Frcmmer, R.W. Vavra, Jr., and V.A. Loy. 1974. Field evaluation of black fly control aerial applications. Mosquito News 34(3): 330-332. Cariaso, B.L. 1962. The ecology of Simulium (Simuliidae, Diptera) aquatic stages. Philippine Agr. 46(5): 369-377. Carlsson, G. 1967. Environmental factors influencing biackfly populations. Bull. World Health Organ. 37: 139-150. Carlsson, G. 1969. Some Simuliidae (Diptera) from Southern Spain. Entomol. MEDD 37(3): 202-206. Chance, M.M. 1970. The functional morphology of the mouthparts of biackfly larvae (Diptera: Simuliidae). Quaest. Entomol. 6(2): 245-284. Chut tor, F.M. 197?. Notes on the biology of South African Simuliidae, particularly Simulium (Eusimuliu.it) nigvitavse Coquillett. Newsletter Limnol. See. Southern Africa 18: 10-18. Cook, R.S. 1971. Leuooaytozoon Danilewsky 1890. Pages 291-299 in J.W. Davis, R.C. Anderson, L. Karstad and D.O. Trainer, eds. Infections and parasitic diseases of wild birds. Iowa State Univ. Press, Ames. Cooke, C.W. 1939. Scenery of Florida, interpreted by a geolopist, Bull. Fla. Ceol. Surv. 17. 118 pp.

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-299Coquillett, D.W. 1898. The buf falo-gnats , or black-flies, of the United States. U.S. Dep. Agr. Div. Entomol. Bull. 10: 66-69. Coquillett, D.W. 1902. New Diptera from North America. Proc. U.S. Nat. Mus. 25: 83-126. Corredor, D. 1975. The black flies of Washington (Diptera: Simuliidae) M.S. Thesis, Washington State Univ. 94 pp. Coscaron, S., and P. Wygodzinsky. 1973. Notas sobre Simulidos Neotropicales. II. Sobre Simuliwn (Psaroniocompsa) opalinifrons (Enderlein) y notas sobre el subgcnero (Insecta, Diptera). Physis SECC. C. 32(84): 161-172. Craig, D.A. 1974. The labrum and cephalic fans of larval Simuliidae (Diptera: Nematocera). Can. J. Zoo]. 52(1): 133-159. Craig, D.A. 1975. The larvae of Tahitian Simuliidae (Diptera: Nematocera). J. Med. Entomol. 12(4): 463-476. Crans, W.J., and L.G. McCuiston. 1970a. A checklist cf the black flies of New Jersey (Diptera: Simuliidae). Mosquito News 30(4"> : 654655. Crans, W.J., and L.G. McCuiston. 1970b. The current status of black fly investigations in New Jersey. N.J. Mosquito Exterm. Ass. Proc. 57: 103-105. Crosskey, R.W. 1957. The Simuliidae (Diptera) of northern Nigeria. Bull. Entomol. Res. 48(1): 59-74. Crosskey, R.W. 1969. A reclassification of the Simuliidae (Diptera) of Africa and its islands. Bull. Brit. Mus. (Natur. Hist.) Entomol. Suppl. 14. 195 pp. Crosskey, R.W. 1973. Family Simuliidae. Pages 423-430 in M.D. Delfinado and D.E. Hardy, eds . A catalogue of the Diptera of the Oriental Region. Vol. 1. Suborder Nematocera. Univ. Press of Hawaii, Honolulu. 618 pp. Crosskey, R.W., and B.V. Peterson. 1972. The Simuliidae described by N. Baranov and their types (Diptera). Bull. Brit. Mus. (Natur. Hist.) Entomol. 27(3): 187-214. Dalmat, H.T. 1955. The black flies (Diptera, Simuliidae) of Guatemala and their role as vectors of onchocerciasis. Smithsonian Miscellaneous Collections 125(1): 425 pp. Datta, M. 1973. New species of black flies (Diptera: Simuliidae) of the subgenera Eusinrulium Roubaud and Gornphostilbia Enderlein from the Darjeeling Area, India. Oriental Insects 7(3): 363-401. Datta, M. 1974. New species of black flies (Diptera: Simuliidae) from the Darjeeling area, India. Oriental Insects 8(4): 457-468.

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-300Datta, M. 1975. Simuliidae (Diptera) from Assam foot-hills, India. Jap. J. Sanit. Zool. 26(1): 31-40. Datta, M., and B. Dasgupta. 1974. Studies on the nocturnal periodicity of six species of black flies (Diptera: Simuliidae) at Uarjeeling, West Bengal. Proc. Indian Acad. Sci. Sec. B 79(4): 147-153. Davies, D.M. 1958. Some parasites of Canadian black flies (Diptera, Simuliidae). Int. Cong. Zool. Proc. 15: 660-661. Davies, D.M. , and B.V. Peterson. 1956. Observations on the mating, feeding, ovarian development, and oviposition of adult black flies (Simuliidae, Diptera). Can. J. Zool. 34: 615-655. Davies, D.M., P,.V. Peterson, and D.M. Wood. 1962. The black flies (Diptera: Simuliidae) of Ontario. Part I. Adult identification and distribution with descriptions of six new species. Proc. Entomol. Soc. Ontario 92: 70-154. Davies, L. 1968. A key to the British species of Simuliidae (Diptera) in the larval, pupal, and adult stages. Freshwater Biol. Ass. Sci. Pub. 24. 125 pp. Davies, L. 1974. Evolution of larval head-fans in Simuliidae (Diptera) as inferred from the structure and biology of Cvozotia crozctensis (Womersley) compared with other genera. Zool. J. Linn. Soc. 55(3): 193-224. Davis, A.N., J.B. Gahan , J. A. Fluno and D.W. Anthony. 1957. Larvicide tests against blackflies in slow-moving streams. Mosquito News 17(4): 261-265. Davis, J.H. 1967. General maps of natural vegetation in Florida. Univ. Fla. Inst. Food Agr. Sci. Agr. Exp. Sta. Circ. S-1.78. DeFoliart, G.R. 1951. The life histories, identification and control of black flies (Diptera: Simuliidae) in the Adirondack Mountains. Ph.D. Dissertation, Cornell Univ. 98 pp. DeFoliart, G.R., and CD. Morris. 1967. A dry ice-baited trap for the collection and field storage of hematophagous Diptera. J. Med. Entomol. 4(3): 360-362. DeFoliart, C.R., and M.R. Rao. 1965. The ornithophilic black fly Simulium meHdionale Riley (Diptera: Simuliidae) feeding on man during autumn. J. Med. Entomol. 2: 84-85. Delfinado, M.D. 1969. Notes on Philippine black flies (Diptera: Simuliidae). J. Med. Entomol. 6(2): 199-207. Delfinado, M.D. 1971. Some Simuliidae and Curtonotidae from the Philippines and the Bismark Islands (Insecta, Diptera). Steenstrunia 1(14): 131-139. (Abstract).

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-301Desser, S.S. 1970. The fine structure of Leucocytozoon simondi. III. The ookinete and mature sporozoite. Can. J. Zool. 48: 641645. Desser, S.S., J.R. Baker, and P.M. Lake. 1970. The fine structure of Leucocytozoon simondi. I. Gametocytogenesis. Can. J. Zoo]. 48: 331-336. Desser, S.S., A.M. Fallis, and P.C.C. Garnham. 1967. Relapses in ducks chronically infected with Leucocytozoon simondi and Pavahaemopvoteus nettionis. Can. J. Zool. 46: 281-285. Desser, S.S., S.B. Mclver, and D. Jcz. 1975. Observations on the role of simuliids and culicids in the transmission of avian and anuran trypanosomes. Int. J. Parasitol. 5(5): 507-510. Dimond, J.B., and W.G. Hart. 1953. Notes or. the blackflies (Simuliidae) of Rhode Island. Mosquito News 13(4): 238-242. Disney, R.H.L. 1969. The timing of adult ecloslon in blackflies (Dipt., Simuliidae) in West Cameroon. Bull. Entomol. Res. 59: 485-503. Disney, R.ll.L. 1971. Association between blackflies (Simuliidae) and prawns (Atyidae), with a discussion of the phcretic habit in simuliids. J. Animal Ecol. 40: 83-92. Disney, R.H.L. 1973. Further observations on some blackflies (Diptera: Simuliidae) associated with mayflies (Ephemeroptera: Baetidae and Heptageniidae) in Cameroon. J. Entomol., London 47: 169-180. Disney, R.H.L. 1975. Speculations regarding the mode of evolution of some remarkable associations between Diptera (Cuterebridae, Simuliidae and Sphaeroceridae) and other arthropods. Entomol. Mon. Mag 110(1311-1321): 67-74. Dove, R.F., and A.B. McKague. 1975. Effects of insect developmental inhibitors on adult emergence of black flies (Diptera: Simuliidae). II. Can. Entomol. 107(11): 1211-1213. Downe, A.E.R., and P.E. Morrison. 1957. Identification of blood meals of black flies (Diptera: Simuliidae) attacking farm animals. Mosquito News 17(1): 37-40. Downes, J. A. 1965. Adaptations of insects in the Arctic. Annu. Rev. Entomol. .10: 257-274. Duke, B.D.L, J. Anderson, and 11. Fuglsang. ] 975. The Onchocerca volvulus transmission potentials and associated patterns of onchocerciasis at four Cameroon Sudan — Savanna villages. Tropenmed. Parasit 26.14 3-154. Dumbleton, L.J. 1963. The classification and distribution of the Simuliidae (Diptera) with particular reference to the genus Austvosimuliwn. New Zeal. J. Sci. 6(3): 320-357.

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-302Dumbleton, L.J. 1972. The genus Austrosimulium Tonnoir (Diptera: Simullidae) with particular reference to the New Zealand fauna. New Zeal. J. Sci. 15(4): 480-584. Dunbar, R.W. 1969. Nine cytological segregates in the Simulium damnosum complex (Diptera: Simuliidae) . World Health Organ. Bull. 40(6)974-979. Dyar, H.G., and R.C. Shannon. 1927. The North American two-winged flies of the family Simuliidae. Proc. U.S. Nat. Mus. 69(10)": 54 pp. Eckhart, P., and R. Snetsinger. 1969. Black flies (Diptera: Simuliidae) of northeastern Pennsylvania. Entomol. Soc. Pennsylvania Melsheimer Entomol. Ser. 4. 7 pp. Edgar, S.A. 1953. A field study of the effect of black fly bites on egg production of laying hens. Poultry Science 32(5): 779-780. Edwards, F.W. 1915. On the British species of Simulium, I. The adults. Bull. Entomol. Res. 6: 23-42. Edwards, F.W. 1920. On the British species of Simulium. II. The early stages; with corrections and additions to part T. Bull. Entomol. Res. 11: 211-246. Elliot, J.M. 1971. Upstream movements of benthic invertebrates in a lake district stream. J. Animal Ecol. 40: 235-252. Emery, W.T. 1914. Morphology and biology of Simulium vittatxm and , its distribution in Kansas. Kansas Univ. Sci. Bull. 8(9)323362. Enderlein, G. 1930. Der heutige stand der klassif ikation der Simuliiden. Arch. Kl. Phylogen. Entomol. 1: 77-97. Enderlein, G. 1936. Simuliologica. Gesell. Naturf. Freunde Sitzber 113-130. Erving, J.M. 1971. Browsing around Florida. Erving Publications, Kissimmee. 174 pp. Ezenwa, A.O. 1973. Mermithid and microsporidan parasitism of blackflies (Diptera, Simuliidae) in the vicinity of Churchhill Falls, Labrador. Can. J. Zool. 51(10): 1109-1111. Fain, A., and P. Elsen. 1973. Notes on the sinmliids of cast Cameroon (Diptera, Simuliidae). (In French). Rev. Zool. Bot. Afr. 87(3)519-554. (Abstract). Fairchild, G.B., and E.A. Barreda. 1945. DDT as a larvicide against Simulium. J. Econ. Entomol. 38: 694-699.

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-303Fallis, A.M., and G.F. Bennett. 1961a. Sporogony of Leucocytozoon and Haemoproteus in simuliids and ceratopogonids and a revised classification of the Haemosporidiida. Can. J. Zool. 39: 215-228. Fallis, A.M., and G.F. Bennett. 1961b. Sporogony of Leucocytozoon and Haemoproteus in simuliids and ceratopogonids and a revised classification of Haemoproteus and other parasites. Mosquito News 21: 21-28. Fallis, A.M., and G.F. Bennett. 1966. On the epizootiology of infections caused by Leucocytozoon simondi in Algonquin Park, Canada. Can J Zool. 44: 101-112. Fallis, A.M., S.S. Desser, and R.A. Khan. 1974. On species of Leucocytozoon. Advances in Parasitol. 12: 1-67. Fallis, A.M., and S.M. Smith. 1964. Ether extracts from birds and C0 9 as attractants for some ornithophilic simuliids. Can. J Zool 42: 723-730. Field, G., R.J. Duplessis, and A. P. Breton. 1967. Progress report on laboratory rearing of black flies (Diptera: Simuliidae) . J. Med Entomol. 4(3): 304-305. Field G., and W. Low. 1961. Dimorphism in simuliid pupae. Ann. Entomol. Soc. Atner. 54: 617-618. Forbes, S.A. 1912. On black flies and buffalo gnats {Simulium) as possible carriers of pellagra in Illinois. Pages 21-55 in 111 State Entomol. Rep. 27. — Forrester, D.J., L.T. Hon, L.E. Williams, Jr., and D.H. Austin. 1974 Blood protozoa of wild turkeys in Florida. J. Protozool. 21(4)494-497. Fredeen, F.J.H. 1973. Black flies. Can. Dep. Agr. Pub. 1499. 19 pp. Fredeen, F.J.H. 1974. Tests with single injections of methoxychlor black fly (Diptera: Simuliidae) larvicides in large rivers Can Entomol. 106: 285-305. Fredeen, F.J.H. 1975. Effects of a single injection of methoxychlor black-fly larvicide on insect larvae in a 161-km (100-mile) section of the North Saskatchewan River. Can. Entomol. 107: 807-817. Fredeen, F.J.H., J.G. Saha , and M.H. Balba. 1975. Residues of Methoxychlor and other chlorinated hydrocarbons in water, sand, and selected fauna following injections of Methoxychlor black fly larvicide into the Saskatchewan River, 1972. Pestic Monit J 8(4): 241-246. ' ' Fredeen, F.J.H., J.G. Saha, and L.M. Rover. 1971. Residues of DDT, DDE, and DDD in fish in the Saskatchewan River after using DDT as a blackfly larvicide for twenty years. J. Fish. Res Bd °Cnn 28(1): 105-1C9.

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-314Tarshis, I.B. 1968. Collecting and rearing black flies. Ann. Entomol. Soc. Amer. 61(5): 1072-1083. Tarshis, I.B. 1971. Individual black f]y rearing cylinders (Diptera: Simuliidae). Ann. Entomol. Soc. Amer. 64(5): 1192-1193. Tarshis, I.B. 1972. The feeding of some ornithophilic black flies (Diptera: Simuliidae) in the laboratory and their role in the transmission of Leucooytozoon simondi. Ann. Entomol. Soc. Amer. 65(4): 842-848. Tarshis, I.B. 1973. Studies on the collection, rearing, and biology of the blackfly (Cnephia ornithophilic). U.S. Dep. Interior Fish Wildlife Serv. Spec. Sci. Rep. 165. 16 pp. Tarshis, I.B., and T.R. Adkins, Jr. 1971. Equipment for transporting live black fly larvae (Diptera: Simuliidae). Ann. Entomol. Soc. Amer. 64(5): 1194-1195. Tarshis, I.B., and W. Neil. 1970. Mass movement of black fly larvae on silken threads (Diptera: Simuliidae). Ann. Entomol. Soc. Amer. 63(2): 607-610. Tarshis, I.B., and J.H. Stuht. 1970. Two species of Simuliidae (Diptera), Cnephia ovnithophilia and Prosimuliurn vernale, from Maryland. Ann. Entomol. Soc. Amer. 63(2): 587-590. Thorsteinson, A. J. , G.K. Bracken, and W. Hanec. 1965. The orientation behavior of horse flies and deer flies (Tabanidae Diptera). III. The use of traps in the study of orientation of tabanids in the field. Entomol. Exp. and Appl. 8: 189-192. Townsend, C.H. 1891. A new Simuliwn from southern New Mexico. Psyche 6: 106-107. Townsend, C.H. 1897. Diptera from the lower Rio Grande or Tamaulipan region of Texas. I. J. N.Y. Entomol. Soc. 5: 171-178. Townsend, L.H., Jr., and E.C. Turner, Jr. 1976. Field evaluation of several chemicals against ear-feeding black fly pests of horses in Virginia. Mosquito News 36(2): 182-186. Travis, B.V., M.H. Goodwin, and W.E. Gambrell. 1939. Preliminary note on the occurrence of Leucooytozoon smithi Laveran and Lucet (1905) in turkeys in the southeastern United States. J. Parasitol. 25: 274. Travis, B.V., D.C. Kurtak, L.C. Rimando. 1970. Tests (1969) with black fly larvicides. N.J. Mosquito Exterm. Ass. Proc. 57: 106-110. Travis, B.V. and R.M. Labadan. 1967a. Black flies. Pages 323-34 3 in Arthropods of medical importance in Asia and the European U.S.S.R. Part II. Tech. Rep. 67-65-ES. U.S., Army Natick Laboratories, Natick, Mass.

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-315Travis, B.V., and R.M. Labadan. 1967b. Black flics. Pages 217-244 In Arthropods of Medical Importance in Latin America. Part II. Tech. Rep. 68-30-ES. U.S. Army Natick Laboratories, Natick, Mass. Travis, B.V., R.M. Labadan, and H.H. Lee. 1968. Black flies. Pages 121-127 in Arthropods in Medical Importance in Australia and the Pacific Islands. Tech. Rep. 68-61-ES. U.S. Army Natick Laboratories, Natick, Mass. Travis, B.V. , H.H. Lee, and R.M. Labadan. 1969. Black flies. Pages 111-143 in Arthropods of medical importance in America north of Mexico. Tech. Rep. 69-2-ES. U.S. Army Natick Laboratories, Natick, Mass. Travis, B.V., C.E. Mendoza, and R.M. Labadan. 1967. Black flies. Pages 359-396 in Arthropods of medical Importance in Africa. Part II. Tech. Rep. 67-55-ES. U.S. Army Natick Laboratories, Natick, Mass. Travis, B.V. and M.V. Vargas. 1970. Bionomics of black flies in Costa Rica. II. An ecological consideration. N.J. Mosquito Exterm. Ass. Proc. 57: 111-112. Travis, B.V. , M.V. Vargas, and J.C. Swart zwelder. 1974. Bionomics of black flies (Diptera: Simuliidae) in Costa Rica. I. Species biting man, with an epidemiological summary for the Western Hemisphere. Rev. Biol. Trop., San Jose, Costa Rica 22(1): 187-200. Tucker, E.S. 1920. Occurrences of black flies in Louisiana during recent years. Trans. Kansas Acad. Sci. 29: 65-75. Twinn, C.R. 1936. The blackflies of eastern Canada (Simuliidae, Diptera). Can. J. Res., D 14: 97-150. Twinn, C.R. 1938. Blackflies from Utah and Idaho, with descriptions of new species (Simuliidae, Diptera). Can. Entomol. 70: 48-55. Uemoto, K. , 0. Onishi, and T. Orii. 1973. Revision of the genus Prosimdiwn Roubaud (Diptera, Simuliidae) of Japan. 1. Hirtipes group in the subgenus Prosimulium. Jap. J. Sanit. Zool. 24(1): 27-46. Underhill, G.W. 1939. Two simuliids found feeding on turkeys in Virginia. J, Econ. Entomol. 32: 765-768. Underhill, G.W. 1944. Blackflies found feeding on turkeys in Virginia, (Simulivn nigropavvwn Twinn and Sinrulium slovsoiaa Dyar and Shannon). Va. Agr. Exp. Sta. Tech. Bull. 94. 32 pp. U.S.D.A. 1976. Black flies. U.S. Dep. Agr. Coop. Plant Pest Rep. 1(17): 192.

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-316 Ussova, Z.V. 1961. Files of the Karelia and the Murmansk Region (Diptera, Slmuliidae) . Academy of Sciences of the U.S.S.R. 268 pp. (Israel Program for Scientific Translations. Jerusalem, 1964.) Vajime, C.G., and R.W. Dunbar. 1975. Chromosomal identification of eight species of the subgenus Edwards el turn near and including Simulium ( 'Edwards ellum) damnosum Theobald (Diptera: Simuliidae). Tropenmed. Parasitol. 26: 111-138. Van Soraeren, G.R.C. 1944. Some records of Simuliidae in Abyssinia and British Somaliland. Bull. Entomol. Res. 35: 113-114. Volkmar, F. 1929. Observations of Lcueocytozoon smithi; with notes on Leuaocytozoa in other poultry. J. Parasitol. 16: 24-28. Vulcano, M.A. 1967. Family Simuliidae. Pages 1-44 in A catalogue of the Diptera of the Americas south cf the United States. 16. Dep . Zool. Seer. Agr., Sao Paulo, Brazil. Walker, F. 1848. List of specimens of dipterous insects. List Dipt. Brit. Mus. 1: 232 pp. Wallace, R.R. , A.S. West, A.E.R. Downe, and H.B.N. Hynes. 1973. The effects of experimental blackfly (Diptera: Simuliidae) larviciding with Abate, Dursban, and Methoxychlor on stream invertebrates. Can. Entomol. 105(6): 817-831. Wehr, E.E. 1953. Report of the chief of the Bureau of Animal Industry. U.S. Dep. Agr. Annu. Rep. 1953: 83. Wehr, E.E. 1962. Studies on Leucocytozoonosis of turkeys, with notes t on schizogony, transmission, and control of Leucocytozoon smithi. Avian Diseases 6: 195-210. Welch, H.E. 1964. Mermithid parasites of blackflies. World Health Organ. Bull. 31: 857-863. Wellington, W.G. 1974. Black-fly activity during cumulus-induced pressure fluctuations. Environ. Entomol. 3(2): 351-353. Wenk, P. 1965. Uber die biologic blutsaugender Simuliiden (Diptera). III. Kopulation, blutsaugen und eiablage von Boophihora erythroeephala de Geer im laboratorium. Tropenmed. Parasitol. 16(2): 207-226. Wenk, P. and J.N. Raybould. 1972. Mating, blood-feeding and oviposition of the "Kibwezi" form of Simulium damnosum Theobald (Diptera: Simuliidae) in the laboratory. World Health Organ. Bull. 47: 627-634. West, A.S., W.F. Baldwin, and J, Gomery. 1968. A radioisotopic-sticky trap-autoradiographic technique for studying the dispersal of black flies. World Health Organ. Vector Biology Control 71.280: 18 pp.

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-317Wirth, W.W., and A. Stone. 1956. Family Slmuliidae. Pages 401-406 In R.L. Usinger, ed. Aquatic insects of California. Univ. of Cali77 Press, Berkeley. 508 pp. Wolfe, L.S., and D.G. Peterson. 1959. Black flies (Diptera: Simuliidae) of the forests of Quebec. Can. J. Zool . 37: 137-159. Wolfe, L.S., and D.G. Peterson. 1960. Diurnal behavior and biting habits of black flies (Diptera: Simuliidae) in the forests of Quebec. Can. J. Zool. 38: 489-497. Wood, D.M., B.V. Peterson, D.M. Davies, and H. Gyorkos. 1963. The black flies (Diptera: Simuliidae) of Ontario. Part II. Larval identification with descriptions and illustrations. Proc. Entomol. Soc. Ontario 19: 99-129. Wood, R. , and E.A. Fernald. 1974. The new Florida atlas. Patterns of the sunshine state. Trend Publications Inc. Rose Printing Co., Tallahassee. 119 pp. World Health Organization. 1971. IV. Blackflies in the Americas. World Health Organ. Vector Biology Control 71.283. 23 pp. Wu, Y.F. 1930. A contribution to the biology of Simuliiun (Diptera). Papers Mich. Acad. Sci. 13: 543-599. Wygodzinsky, P. 1971. Descriptions and redescriptions of species of the blackfly genus Sinruliwn from the Northern Andes (Simuliidae, Diptera). Amer. Mus. Novitates 2447. 38 pp. .Wygodzinsky, P. 1973. On a species of Simuliur; (Ectenmaspis) from the Northeastern United States (Diptera). N.Y. Entomol Soc J 81(1): 10-12. Wygodzinsky, P., and S. Coscaron. 1970. A new species group of the blackfly genus Sinruliwn from western South America (Simuliidae, Diptera, Insecta) . Amer. Mus. Novitates 2< / ;33. 20 pp. Wygodzinsky, P., and S. Coscaron. 1973. A review of the Meso American and South American black flies of the tribe Prosirouliini (Simuliinae, Simuliidae). Bull. Amer. Mus. Natur. Hist. 151(2): 199 pp. Wygodzinsky, P., and A.D. Najera. 1970. Un nucvo genero de simulido de la Republica Mexicana (Diptera: Simuliidae). Pev. Invest. Salud. Pub., Mexico, 30: 83-110. Yang, Y.J., and D.M. Davies. 1974. The saliva of adult blackflies (Simuliidae: Diptera). Can. J. Zool. 52(6): 749-753, Zetterstedt, J.W. 1838. S. vittata. Insecta Lapponica (1838-1840)803. Zivkovic, V. 1971. Black flies (Diptera, Simuliidae) of the Danube ii Yugoslavia. Acta Vet. 21(5): 225-236. (Abstract).

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-31 J Zwick, H. 1974. Faunistical, ecological and taxonomlc studies on Simullidae (Diptera), mainly of the Fulda area. (In German). Senckenb. Na'curforsch. Ges. 533: 116 pp. Abh,

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APPENDIX COLLECTION SITE NAMES AND LOCATIONS Alachua Co. 1) Sandy Hatchet Creek. 3.4 km north of Highway 24 on State Road (SR) 225. R 20 E-T 9 S, Section 2, SE quadrant. 2) Blues Creek. 1.6 km north of SR 222 at the 1FAS Horticulture Unit. R 19 E-T 9 S, Section 9, SW quadrant. 3) Devil's Millhopper. 2.4 km north of SR 222 on SR 232. R 19 E-T 9 S, Section 15, 5E quadrant. 4) Tributary to Hogtown Creek. .8 km south of the Devil's Millhopper on SR 232. il 19 E--T 9 S, on the border of Sections 22 and 23. 5) Gainesville. (Adult black fly captured biting a human — D. Young.) 6) Tributary of Hatchet Creek. 5.6 km northeast of the SR 225/Highway 24 junction, on Highway 24. R 21 E-T 9 S, Section 6, NE quadrant. 7) Little Hatchet Creek. 1.6 km northeast of the SR 232/Highway 24 < ' junction, on Highway 24. R 20 E-T 9 S, Section 23, NE quadrant. 8) Tributary to the Santa Fe River. .3 km north of the SR 225/340 junction, on SR 225. R 20 E-T 8 S, Section 15, NE quadrant. 9) Tributary to the Santa Fe River. 1.3 km northeast of the SR 225/ 340 junction, off SR 225 in the woods. R 20 E-T 9 S, Section 14, NW quadrant. 10) Tributary to the Santa Fe River. 1 km west of the SR 225/340 junction, on SR 340. R 20 E-T 8 S, Section 15, NW quadrant. 11) Stream to Newnan's Lake. 1 . 2 km north of SR 225/Highway 20 junction, on SR 225. R 20 E-T 10 S, on the border of Sections 1 and 2. 12) At a cypress dome. 3.2 km northwest of Gainesville off Route 441. (Adult black flies captured from chickens — H. Davis.) 13) Gainesville, Univ. Florida campus. Northeast about 20 ra from the intersection of Museum Road and Center Drive. R 20 E-T 10 S. 14) Tributary to the Santa Fe River. 1.6 km west c»f SR 225. R 20 E-T 8 S, center of Section 27. -319-

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-32015) Hogtown Creek. Gainesville at the junction of University Avenue and 34th Street. R 19 E-T 10 S. 16) Rattlesnake Branch. Gainesville. At the north end of NW 20th Terrace off 8th Avenue. R 20 E-T 10 S. 17) Hatchet Creek. 4 km north of the SR 225/Highwny 24 junction, on Highway 24. R 21 E-T 9 S, Section 7, NW quadrant. 18) Flow to Newnan's Lake. 4.2 km southwest of the SR 232/IIighway 26 junction, on Highway 26. R 20 E-T 10 S, Section 31, SW quadrant. 19) Santa Fe River. SR 225 at Alachua-Bradford County border. R 21 E-T 7 S, Section 32, NE quadrant. 19a) Santa Fe River. SR 235 at Alachua-Bradford County border. R 20 E-T 7 S, Section 18, SE quadrant (Stoneand Snoddy, 1969). 20) Hatchet Creek. Or. Highway 26 1.6 km east of the junction of SR 232/Highway 26. R 21 E-T 9 S, Section 21, SE quadrant. 21) Lochloosa Crock. On Highway 20 6.4 km east of SR 234/Highuay 20 junction. R 22 E-T 10 8. Section 30, NE quadrant. 22) Lochloosa Creek. 6.4 km east of SR 325 and 1.6 km north of Sike's Fish Camp. R 22 E-T 11 S, Section 6, NE quadrant. 23) Tributary to Lochloosa Creek. 4.8 km east of SR 325 and .5 km north of Sike's Fish Camp. .24) Hatchet Creek. Off Highway 26 1.8 km east of the junction of SR 232/Highway 26, .6 km north of the highway in the woods at the Hatchet Creek Preserve. R 21 E-T 9 S, Section 22, SW quadrant. 25) Austin Cary Memorial Forest. 8 km northeast e-f the junction of SR 225/Highway 24, up Highway 24. Adult black flies captured in a Malaise trap by J. Click. 26) Tributary to Hogtown Creek. Gainesville, in the woods east of 34th Street and NW 34th Ave, R 19 E-T 9 S, Section 26, SE quadrant. 27) Tributary to Hogtown Creek. Gainesville, in the woods east of 34th Street and NW 27th Avenue. R 19 E-T 9 S, Section 26, SE quadrant . 28) Gainesville, in the woods south of NW 23rd Avenue and NT' 83rd Street at Santo Fe College. R 19 E-T 9 S, Section 32 NE quadrant. 29) Hogtown Creek. Gainesville, on SR 2 32A 4 km. southeast of Highway 23 at: the Gaineswood Apartments. R 20 E-T 9 S. 32) Hogtown Creek. Gainesville, on SW 20th /Wenu? 3.0 km west of 34th Street. R 19 E-T 10 S, Section 10, SE quadrant.

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-32133) Gainesville, Univ. Florida campus. Stream from Graham and Simpson Hall ponds at intersection of North-South Drive and Museum Road. R 20 E-T 10 S. 34) Gainesville, Univ. Florida campus. Stream near Veterinary Entomology Building at the bottom of the hill on Newell Drive. R 20 E-T 10 S. 36) Headwaters of Hatchet Creek. Off SR 225 .8 km due west, across from NE 73 Avenue. R 20 E-T 9 S, Section 11, SW quadrant. 38) Lacrosse, Jamnback and Stone, 1957. 39) Gainesville, at the spillway just east of the Patuxent Wildlife Research Center on Highway 26 2.2 km west of SR 225. R 20 E-T 10 S. Section 2, west center. Baker Co. 41) Turkey Creek. 1.6 km south of Route 90 on SR 121 at MacClenny . R 22 E-T 3 S, Section 5, NE quadrant. Bay Co. 40) Juniper Creek. 4.8 km north of Highway 20 on Route 231. R 12 W-T 1 N, Section 14, west center. Bradford Co. 42) Prevatt Creek. Near Starke, 5.6 km north of the junction of Highway 18 and Route 301, on Route 301. R 22 E-T 7 S, Section 6, SE quadrant. 43) Double Run Creek. 4 km southeast of Hampton on Highway 18. R 22 E-T 7 S, Section 27, SW quadrant. 44) Water Oak Creek. 3.5 km north of the Highway 16/Route 301 junction, on Route 301, north cf Starke. R 22 E-T 6 S, Section 10, SW quadrant . 45) Sampson Creek. 8 km west of the Highway 18/Route 301 junction, on Highway 18, west of Graham. R 21 E-T 7 S, Section 29, NW quadrant. 46) Drainage flow to Double Run Creek. 4 km southeast of Hampton on Highway 18. R 22 E-T 7 S, Section 27, SW quadrant. 47) 02ust.ee Creek. In Lawtey .4 km north of the SR 225/Route 301 junction, on Route 301. R 22 E-T 5 S, Section 23, SE quadrant. 48) Small flow west of Brooker on Highway 18 at the junction with SR 237. R 29 E-T 7 S, Section 1, SE quadrant. 49) Juniper Creek. 9.7 km west of Clarkesville on Highway 20. R 21 W-T 1 S, Section 1, west center.

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-32250) Chipola River. 2.4 km west of SR 275 on Highway 20. R 9 W-T 1 S, Section 6, NE quadrant. 51) Apalachicola River. 3.0 km east of Blountstown on Highway 20. R 8 W-T 1 N, Section 35, NE quadrant. 52) Four Mile Creek. 1.4 km north of the Highway 73/Highway 20 junction, on Highway 73. R 9 W-T 1 N, Section 31, NW quadrant. 53) Blountstown. J. Yelvington collected S. mevidionale adults on 10 May 19 73 from young chickens. R 8 W-T 1 N. Clay Co. 55) Gold Head Branch. At Gold Head Branch State Park 9.7 km northeast of SR 100/Mighway 21 junction on Highway 21. R 23 E-T 8 S, Section 1, NE quadrant. 56) Tributary to Ates Creek. 8.9 km northeast of Gold Head Branch State Park on Highway 21. R 24 E-T 7 S, Section 3, SW quadrant. 57) South Prong Black Creek. 4 km south of Highway 16 on Highway 21. R 24 E-T 6 S, Section 27, SW quadrant. Columbia Co. 58) Santa Fe River. At Oleno State Park 3.2 km east of Route 441. R 17 E-T 7 S, Section 2, NE quadrant. 59) Ichetucknee River. On Route 27 at the Columbia/Suwanee County border. R 15 E-T 6 S, Section 23, NW quadrant. 60) Santa Fe River. On Route 27 at the Alachua/Columbia County border north of High Springs. R 17 E-T 7 S, Sect ten 32, NE quadrant. (Young, undetermined S. (PhosterodOTOs) spp. larvae collected.) 61) Santa Fe River. 2.4 km south of SR 138 at the Gilchrist/Columbia County line. R 16 E-T 7 S, Section 36, SW quadrant. (Young, undetermined S. (Phosterodoros) spp. larvae collected.) 62) Santa Fe River. 2.4 km south of SR 138 at the Gilchrist/Columbia County line, side flow in the woods. R 16 E-T 7 S, Section 36, SW quadrant. Dade Co. 63) Biscayne Bay, Florida. Mrs. A.T. Slcsson, black fly adults. Desoto Co. 64) Prairie Creek. 2.4 km northwest of the Charlotte/Desoto County line on Highway 31. R 25 E-T 39 S, Section 26, SE quadrant. 65) Joshua Creek. On Highway 31 .2 Ian north of the Highway 31/SR 760 junction, southeast of Arcadia. R 25 E-T 38 S, Section 16, NW quadrant. 66) Buzzard Roost Branch. On Highway 70 13 km northwest of Arcadia, R 23 E-T 37 S, Section 10, SW quadrant.

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-32 3Dixic Co. 67) Tributary to the Steinhatchee River. 23 km northwest of Cross City on Route98. R 10 E-T 8 S, Section 33, NE quadrant. 68) On Route 98 within 1.6 km northwest of Cross City. R 12 E-T 10 S, Section 5, SE quadrant. 69) Pumpkin Swamp Creek. 13 km northwest of Cross City on Route 98. R 11 E-T 9 S, Section 20, SW quadrant. Duval Co. 70) Yellow Water Creek. 9.7 km northeast of SR 228/Route 301 junction on SR 228. R 24 E-T 3 S, Section 20, NE quadrant. 71) Jacksonville. Mrs. A.T. Slosson, black fly adult. 72) 6.4 km west of the SR 115c/Route 90 junction on Route 90. R 24 E-T 2 S, Section 20, SE quadrant. Escambia Co. 73) Canoe Creek. 13.7 km into Escambia Co. going west on Highway 4. R 31 W-T 5 N, Section 8, NW quadrant. 74) Pine Barrens Creek. 4.8 km south of the Highway 4/Highway 99 junction, on Highway 99. R 32 W-T 5 N, Section 28, NE quadrant. 75) McDavid Creek. 17.7 km south of the Highway 97 and Highway 99 junction, on Highway 99. R 32 W-T 3 N, Section 33, NE quadrant. Flagler Co. 77) Little Haw Creek. 3.2 km east of the .Volusia/Flagler County line on SR 305. R 29 E-T 14 S, Section 5, NE quadrant. Franklin Co. 80) Hickory Landing State Park. 3.2 km west of Highway 65. R 8 W-T 6 S, Section 2, NW quadrant. G.B. Fairchild, black fly adults. Gadsden Co. 81) Tributary to the Ochlockonee River. 1.1 km west of Concord on Highway 12. R 1 W-T 3 N, Section 18, SW quadrant. 82) Holman Branch Quincy Creek. Highway 12 at the eastern Quincy city limits. R 3 W-T 2 N, Section 8, NW quadrant. 83) Hurricane Creek. 1.1 km southeast of the Highway 12/SR 270 junction on SR 270. R 3 W-T 2 N, Section 23, SW quadrant. 85) Crooked Creek. SR 270. .4 km east of the Liberty/Gadsden County line. R 6 W-T 2 N, Section 7, SW quadrant. 86) Shepard's Mill. 1.1 km east of the SR 274/Highway 12 junction, on Highway 12, east of Greensboro. R 5 W-T 2 N, Section 15, SE quadrant.

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-32487) Tributary to the Ochlockonee River. 1.6 km west of Concord on Highway 12. R 2 W-T 3 N, Section 13, SE quadrant. 88) Chattahoochee. Power station dam at the State hospital on Route 90. R 6 W-T 4 N. 89) Chattahoochee. Mosquito Creek, below second power station dam at the State hospital on Route 90. R 6 W-T 4 N. 90) Tributary to the Ochlockonee River. 1.6 km northeast of Havana on Highway 12. R 2 W-T 3 N, Section 25, NW quadrant. 91) Shaw Creek. 1.6 km east of the Leon/Gadsden County line on Highway 12. R 1 W-T 3 N, Section 15, NE quadrant. Glades Co. 95) FisheaLing Creek. West of Route 27 at Palmdale. R 29, 30 E-T 40, 41 S. Gulf Co. 97) St. Joseph State Park. W.W. Wirth, black fly adult. 98) Beacon Hill. W.W. Wirth, black fly adult. Hamilton Co. 99) Alapaha River. 3 km northeast of the Interstate 75/Highway 6 junction, on Highway 6. R 13 E-T 2 N, Section 32, SE quadrant. 100) Sugar Creek. 5.6 km south of the Interstate 75/Route 129 junction, on Route 129. R 14 E-T 1 S, Section 17, SE quadrant. 101) Small flow .32 km northwest of the Interstate 75/Route 129 junction on Interstate 75. R 14 E-T 1 N. Hardee Co. 102) Troublesome Creek. 6.4 km west of Zolfo Springs on Highway 64. R 24 E-T 34 S, Section 36, NE quadrant. 103) Thompson Branch. 1.6 km south of Wachula on Route 17. R 25 E-T 34 S, Section 15, NW quadrant. 104) Hog Branch. 4 km north of Wachula on Route 17. R 25 E-T 33 S, Section 21, SW quadrant. Hendry Co. 106) Banana Branch. 6.4 km southwest of Labelle on Highway 80. R 28 E-T 43 S, Section 27, NE quadrant. Highlands Co. 108) Little Charley Bowlegs Creek. At Highlands Hammock State Park, 8 km west of Route 27. R 28 E-T 34 S, Section 31, south center. 109) Small flow at Highlands Hammock State Park, restricted area. 5.6 km west of Route 27. R 28 E-T 34 S, Section 33, SW quadrant.

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•325110) Archbold Biological Station near Lake Placid. Highway 70/Route 27 junction. R 30 E-T 38 S. Hillsborough Co. 111) Turkey Creek. 6.4 km west of Highway 39/60 junction on Highway 60. R 21 E-T 29 S, Section 22, SE quadrant. 112) Hillsborough County. J. Cross, adult black fly. 113) Blackwater Creek. 3.6 km south of the Pasco/Hillsborough County line on Highway 39. R 22 E-T 27 S, Section 18, NW quadrant. 114) Trout Creek. 14.5 km north of Sr 582/SR 581 junction on SR 581. R 19 E-T 27 S, Section 13, NW quadrant. Holmes Co. 116) Blue Creek. At the junction of Highway 81 and Route 90 in Ponce DeLeon. R 17 W-T 3 N, Section 28, NE quadrant. 117) Little Reedy Creek. 2.4 km northeast of Ponce De Leon on Route 90. R 17 W-T 3 N, Section 23, SE quadrant. 118) Camp Branch. 2.7 km west of the Washington/Holmes County line on Route 90, east of Bonifay. R 14 W-T 3 N, Section 4, NW quadrant. 119) Cum Creek. 3.2 km west of Bonifay on Route 90. R 15 W-T 4 N, Section 34, SE quadrant. Jackson Co. 120) Florida Caverns State Park. W.W. Wirth, adult black flies R 10 W-T 5 N. 123) Alford. D.E. Hardy, adult black fly. R 12 W-T 3,4 N. Jefferson Co. 124) 3.2 km east of Capps on Route 27. R 4 E-T 1 S. 125) Burnt Mill Creek. 1.5 km east of the Leon/Jefferson County line on Route 27. R 3 E-T 1 S, Section 7, SE quadrant. Lafayette Co. 129) Kettle Creek. 1.6 km north of the Dixie/Lafayette County line on Highway 51. R 10 E-T 7 S, Section 27, SW quadrant. 130) Kettle Creek (a northern fork). 2.2 km north of the nixie/Lafayette County line on Highway 51. R 10 E-T 7 S, Section 27, center. Lake Co. 170) Blackwater Creek. 18.5 km east of Eustls on Highway 44. R 28 E-T 18 S, Section 35, SW quadrant. Leon Co. 131) Tall Timbers Research Station, .stream to Gannet Pond. 6.4 km west of Route 319 and just south of Highway 12. R 1. F.-T 3 N.

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-326132) Tall Timbers Research Station, stream from Gannet Pond. 6.4 km west of Route 319, just south of Highway 12 and north of Lake Iamonia. R 1 E-T 3 N. 134) Black Creek. 5.6 km west of Jefferson/Leon County line on Route 90. R 2 E-T 1 N, Section 11, NE quadrant. Levy Co. 135) Otter Creek. 1 . 2 km southwest of the Route 98/Highway 24 junction, on Highway 24. R 15 E-T 13 S, Section 26, SW quadrant. 136) Waccasassa River. 6.4 km southeast of the Route 98/Highway 24 junction, on Route 98. R 16 E-T 14 S, Section 17, NW quadrant. 137) Waccasassa River. 4.8 km northeast of the Route 98/Highway 24 junction, on Highway 24. R 16 E-T 13 S, Section 17, NW quadrant. 138) Side flow to the Waccasassa River. 4.8 km northeast of the Route 98/Highway 24 junction, on Highway 24. R 16 E-T 13 S, Section 17, NW quadrant. 139) North Prong Tenmile Creek. .2 km southeast of the Route 98/SR 336 junction, on SR 336. R 16 E-T 15 S, Section 24, east center. 140) Wekiva River. 1 . 8 km northeast of Route 98 on SR 326. R 16 E-T 14 S, Section 16, SE quadrant. 141) Stream at Gulf Hammock. On Route 98 .4 km southeast of SR 326. R 16 E-T 14 S, Section 21, SW quadrant. Liberty Co. 142) Tributary to Telogia Creek. On Highway 20 1.6 km west of Hosford. R 6 W-T 1 S, Section 14, NW quadrant. 143) Camel Lake State Park. G.B. Fairchiid, adult black flies. R 8 W-T 2 S. 144) West Minnow Branch". On SR 271 3.7 km south of Highway 12. R 6 W-T 1 N, Section 19, NE quadrant. 145) Apalachicola River. At Richbourg Landing just off SR 333 2.4 km southwest of the SR 333/Highway 12 junction. R 8 W-T 2 S, Section 16, NE quadrant. 146) Telogia Creek. On SR 271 5.6 km south of Highway 12. R 6 W-T 1 N, Section 30, SE quadrant. 147) B]ue Creek. 4.8 km east of Hosford on Highway 20. R 5 W-T 1 S, Section 21, NW quadrant. 148) Rock Creek. At Torreya State Park north of SR 271. R 7 W-T 2 N. |149) Telogia Creek. 6.4 km west of SR 271 on Highway 20. R 7 W-T 1 S, Section 3, NW quadrant.

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-327150) Upper Sweetwater Creek. 8 km northwest of Highway 12 on SR 270. R 7 W-T 2 N, Section 31, east center. 154) Torreya State Park. F.S. Blanton, H.V. Weems, W.W. Wirth, adult black flies. R 7 W-T 2 N. Madison Co. 151) Withlacoochee River. 12 km on Highway 6 east of the Route 90/Highway 6 junction, R 11 E-T 1 N, Section 17, SW quadrant 152) Withlacoochee Rivt r. W. Beck, black fly Ian 153) Tributary to the Aucilla R iver. On Route 27 3.2 km southeast of the Jefferson/Madison County line. R 5 E-T 2 S, Sect quadrant. ion 2, NE Manatee Co. 155) Wingate Creek. 8.8 Highway 64. R 22 E-T 35 S, Sect west of the Hardee/Manatee County line ion 6, SE quadrant 156) Myakks River. 5.6 km west of the Hardee/Manatee County lii Highway 64. R 22 E-T 35 S, Section 4, SI quadrant 157) Manatee River. 8.9 km east of the SR 675/Highway 64 junctic Highway 64. R 21 E-T 34 S, Section 33, SE quadrant. Marion Co , 158) A small creek on Highway 40. W. Beck, black fly 1 arvae. Nassau Co. 159) Thomas Creek. 12.9 km northeast of the Duval/Nassau County line on Route 301. R 24 E-T 1 N, Section 28, NE quadrant. 160) Small flow 21 km northeast of the Duval/Nassau County line on Route 301, south of Callahan. R 24 E-T 1 N, Section 1 SW quadrant 161) Tributary to Boggy Creek. 5.6 km northeast of Route 301 on Mussel White Road. R 25 E-T 3 N, Sect ion 34, SW quadrant 162) Spell Creek. 8 km northeast of Route 301 on Mussel White Road. R 25 E-T 3 N, Section 27, NE quadrant , Okaloosa Co. 163) Blackwater River. 12.1 km northwest of Route 90 on Highway 4 R 25 W-T 4 N, Secti SW quadrant 164) Blackwater River. 4.8 km west of Holt. R 25 W-T 3 N, Section 31 NW quadrant. |165) Muddy Brook. 5.6 km northwest of the Blackwater River 4. R 25 W-T 4 N, Section IS, NW quadrant. on Highway Il66) Pinev Woods Creek. 3.2 km east of Crestvi W-T 3 N, Section 14, SW quadrant ew on Route 90. R 23

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-328167) Brackin Road Spillway, Crestview. Just north of Route 90. R 23 W-T 3 N. 168) Tributary to the Blackwater River. 16 km west of the Route 90/ Highway 4 junction and 4 km north of Route 90 in the woods. R 25 W-T 3 N. 224) Destin. Adult black fly specimen from A.J. Rogers. R 22 , 23 W-T 2 S. Orange. Co. 169) Tributary to the Econlockhatchee River. On Highway 50 .2 km east of SR 420. R 31 E-T 22 S, Section 23, NE quadrant. 171) Big Econlockhatchee River, Orlando. H.K. Gouck, black fly pupae. 174) Little Econlockhatchee River and Orlando. H.K. Gouck, W.V. King and K.M. Sotnmerman, black fly immatures and adults. 176) Narcussee Swamp, Pine Castle. Black fly pupa, collector unknown. Osceola Co. 179) Kissimee. A.L. Melander, black fly adults. Pasco Co. 177) Stream from a lake. 6.4 km north of Highway 50 on Route 41. R 18 E-T 24 S, Section 22, NE quadrant. 178) Hudson. D.E. Hardy, black fly adults. Pinellas Co. 180) Dunedin. Black fly adult, collector unknown. Polk Co. 181) Peace River. 1.6 km east of Route 17 on Route 98 at Fort Meade. R 25 E-T 31 S, Section 26, SE quadrant. 182) 6.4 km west of Mulberry on Highway 60. R 24 E-T 30 S. 183) Whidden Creek. 4.8 km south of Route 98 and Fort Meade, on Route 17. R 25 E-T 32 S, Section 10, SW quadrant. Putnam Co. 185) Stream from Lake Melrose. 1.6 km south of the Highway 26 and Highway 21 junction, on Highway 21. R 23 E-T 9 S, Section 19, SE quadrant. 186) 1.6 km east of Crandin on SR 100. R 24 E-T 9 S, Section 9, NE quadrant . 187) Palmetto Branch. 12.9 km west of Grandin on SR 100. R 25 E-T 9 S, Section 15, SW quadrant.

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-329188) Welaka. H.A. Denmark, adult black fly. 189) Little Rice Creek. 19 kin east of Grandin on SR 100. R 26 E-T 9 S, Section 32, NW quadrant. 190) Route 17 3.2 km north of the Cresent City High School. R 27 E-T 12 S. 191) Deep Creek. 19 km south of SR 100, on SR 315. R 24 E-T 11 S, Section 3, NE quadrant. 192) 2.1 km south of Dunn's creek on Route 17. R 26 E-T 11 S, Section 2, SE quadrant. 193) Stream from Mirror Lake at Interlachen. Highway 20 1.2 km east of SR 315. R 24 E-T 1.0 S, Section 14, NW quadrant. Santa Rosa Co. 195) Big Juniper Creek. 3.2 km west of SR 191, on Highway 4. R 27 W-T 4 N, Section 12, SW quadrant. 196) 1.6 km northwest of Highway 87, on Highway 4. R 28 W-T 5 N, Section 28, NW quadrant. 197) Blackwater River. W.W. Wirth, black fly adults. Sarasota Co. 198) Phillippi Creek. W. Beck, black fly larvae. Seminole Co. 202) Howell Creek. 5.2 km east of SR 436 on Red Bug Lake Road. R 30 E-T 21 S, Section 24, east center. 203) Bear Gully Canal. 8 km east of SR 436 on Red Bug Lake Road. R 31 E-T 21 S, Section 19, east center. 204) No specific locale, just Seminole County. E.G. Jay, black fly immatures and adult. Sumter Co. 206) Rutland Creek. On Highway 44 9.6 km west of Interstate 75. R 21 E-T 19 S, Section 10, NE quadrant. Suwanee Co. 207) Suwanee River. Route 27 at Branford. R 14 E-T 6 S, Section 21, NW quadrant. 208) Suwanee River. Route 129 4 km south of Interstate 75 at the Suwanee/Hamilton County line. R 14 E-T 1 S, Section 17, SE quadrant. 209) No specific locale, just Suwanee County. D.J. Taylor, adult black flies.

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-3 30Taylor Co. 210) Fenholloway River. 11 km east of Perry on Route 27. R 8 E-T 4 S, Section 36, NE quadrant. 211) Tributary to the Fenholloway River. 9.4 km east of Perry on Route 27. R 8 E-T A S. 213) Rocky Creek. On Route 27 4.7 km northwest of Perry. R 7 E-T 4 S, Section 4, NE quadrant. 214) California Creek. On Route 27 4.8 km northwest of Salem. R 9 E-T 6 S, Section 29, NE quadrant. 215) Athena Warrior Creek. On Route 27 27 km northwest of Highway 51. R 8 E-T 6 S, Section 11, NE quadrant. Union Co. 216) Turkey Creek. On SR 121 .2 km southwest of Highway 16. R 21 E-T 5 S, Section 17, NW quadrant. 217) Tributary to the Sante Fe River. 3.2 km west of SR 121, on Highway 18. R 19 E-T 6 S, Section 30, NW quadrant. Wakulla Co. 219) Ochlockonee River. W.W. Wirth, adult black fly. Walton Co. 220) Adam's Mill Creek. 4 km north of Route 90, on SR 285. R 21 W-T 3 N, Section 12, NW quadrant. 221) Bay Shoal River. 5.6 km north of Rout;e 90, on SR 285. R 21 W-T 4 N, Section 35, SE quadrant. 222) Bruce Creek. 1.1 km west of SR 183, on SR 280. R 18 W-T 3 N, Section 6, NW quadrant. 223) Panther Creek. 4.8 km west of SR 183 on SR 280. R 18 W-T 3 N, Section 20, SE quadrant.

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BIOGRAPHICAL SKETCH The author was born on February 19, 1947, in Bridgeport, Connecticut. He attended secondary schools in Sumter, South Carolina, and Anchorage, Alaska, and graduated from Anchorage West High in June of 1964. In September 1964 he enrolled as an undergraduate at the. University of Connecticut where he majored in Zoology and minored in Chemistry. While at the University of Connecticut he I iccame a University Scholar and member of the Phi Beta Ka appa and Phi Kappa Phi honor societies. The author was awarded a Bachelor of Arts d egree with highest honors and distinction in Zoology in Jane of 1968. During the same month he beeai graduate studies at Cornell University where h e received a Liberty Hyde Bailey Research Assistantship and majored in Medical Entomolc gy and uiinored in Insect ToxicoL At Cornell the author was a member of the Gamma Alphs Scientific Society. He received a Master of Science dee in Entomology in 1979 and ;gan serving as a commissioned officer in th Biomedical Sciences Corps of the U.S. Air F orce. in 1973 the author was accepted into the Civilian Institutions Pr ogram of the U.S. A ir iorce Institute of Techno! ogy, and entered the Department of EnromoL 'gy and Nematology at the University of Flori< a Ph.D. randiGate. the University of Florida he held tl le position cf President of the Newell Entomological Society for one term. The author is the Entomological Society of America, th e American Mosquito Control Association and the American Registry of Professional Ent< -331

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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.F. Butler, Chairman 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. A.G.B. Fairchild Adjunct Professor Emeritus 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. J-ls~~-—r~y ^^uc4j T.J. Walker -^^ ^A^V Professor of Entomology and Nematology

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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. 4 L iJC-'U-v-^/ D.A. Dame Associate Professor of Entomology 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. D.J. Forrester Associate Professor of Veterinary 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. August, 1976 uk £J> __ College of Agriculture Dean, Graduate School

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UNIVERSITY OF FLORIDA 3 1262 08553 0490