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 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 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. 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. 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 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 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 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 LIST OF FIGURES F1Sure 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 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 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 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 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 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 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. 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 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: Cnephia (Cnephia) peeuarwn (Riley); Simulium (Eusimulium) -1- -2- congareenarum(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 (Phostero- doros) 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 smithi 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 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 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, 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- -4- Parasimulium 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 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 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) - -5- 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 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 -6- scope. 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. -7- vittatum 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 C02~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 -8- 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, 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 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 -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 development. Black fly larvae are usually between 4.5 and 10 mm long when mature, club-shaped with a wider -9- poaterior 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 -10- Disney (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 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 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 -11- flat 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 dis- 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 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 -12- from 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 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 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 -13- eliminate a particulate plug from larval guts than at a higher tempera- ture (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 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. 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. -14- Adults - emergence. Tarshis (1968) reported that pupal develop- ment 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 ex- ceed 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 -15- black 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 fe- males 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 C02 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, 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 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. piatipes flew upside down with their abdomens curved upward, con- tacted females as they flew by overhead, settled to a solid substrate, -17- and 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 re- ported 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 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. 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 -19- of 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 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 (196.3a) 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 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 -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 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. ovnitho- 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. 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 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. -22- Al 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 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. 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 re- sulted 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 epi- demic VEE virus has been isolated from Simuliim species in Colombia and although biologic transmission has not been proven, mechanical trans- mission 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 -23- 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. 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 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 simuliid population levels in streams (Jamnback, 1973). -24- Chemical control. Carestia et al. (1974) reported that aerial ap- plications 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 -25- 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 to 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 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 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 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 -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 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) in- dicates predators are unlikely to serve well as man-manipulated biologi- cal 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 (microspori- dian) 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 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 Coelomyoidium 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 -27- 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 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 -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 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 -29- 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 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 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 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 Brad- 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, 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. 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 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). -31- 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 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 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 northwestern Florida a continental weather pattern predominates whereas a more -32- 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 Plecoptera while 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 caddisflies, simuliid larvae, Plecoptera, Stenonema mayflies, orthocladine chironomids, and Corydalis. 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 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 -33- 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. 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'. -34- 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, 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 respec- tive 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 in- formation 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 -35- also 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 trans- mitted 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 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 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 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. 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 charac- teristic spindle shape (Huff, 1942). Desser et al. (1970) found with L. simondi elongated cells resulted when merozoites developed in leuco- cytes and round cells resulted from invasion of red blood cells. Macro- gametocytes 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 -36- the 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 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 ookinetes 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. smithi invade the liver parenchymal cells of the turkey and grow into hepatic schizonts. Huff (1942) re- ports 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 megalo- 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 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 para- sitemias 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 organ- ism 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, -38- 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. 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 tur- keys' 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 sup- posed 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 -39- 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. smithi 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 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 out- breaks 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% in- cidence 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. 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 iramatures, 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. Imagos could be reared very successfully in this manner from -40- -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 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 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, 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 -42- 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 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 identi- fications 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 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 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 -43- 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. 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 -44- Figure 1. A modified Manitoba trap with a black plastic skirt. -45- 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 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 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 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 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 -46- 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- — ' - > * mm mn^ far****- J^p £***>. Figure 2. Sentinel turkeys in an exposure cage. Figure 3. A blackout box trap in the field. -48- tJPMUL t *-^u&j&ft Figure 4. One view of a ramp trap. *** . * "V c* >v» ■•■-• •■ ■■ •rf ~^^ ^/' JB v j&zt^mm Figure 5. An exposed turkey in the field. -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 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 plastic- covered aquarium or a glass desiccator-type container (Fig. 6) where high humidity was maintained by including a sponge half immersed in a -51- Figure 6. Glass container and paper cartons used for holding black fly adults alive in the laboratory. -52- 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 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 car- tons 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 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. 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 -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% 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. 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- -55- Table 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. -56- rJ J4&- Figure 7. Locations in Florida where black flies have been collected. -57- Table 2. Florida black fly distribution records by county. Species r-l ^ O £< s e V l-«i V ■B 3 o 3 3 3 +4 "XJ CO a 00 •p4 •fl to •^ <3 ts tj a s, fc 00 •gl t to 03j •t» to c 00 00 •^ CO o 44 0 00 00 o s H 3j £ c r^ -p to 44 '*.: -Q c^ si d, 00 IO o >s 8 00 0 « o Ss •^ 00 s 00 SX Q. o S 00 00 "C3 r« "t-3 ■*-a r-i s s +i a ^3 44 a CO to County Sites Co CO CO CO CO CO CO CO CO CO CO CO CO CO CO CO CO ES 3r<3'0'0 ooSc^to cjj ca ssgwscc ^carOOV^b^oaQ^ti'^CaSCaCufi, County Sites cj CO to CO to CO to to to to to to to to to to to to Nassau 4 X X X X X X X Okaloosa 7 X X X X X X X X Orange 4 X X X Osceola 1 X Pasco n X X Pinellas 1 X Polk 3 X X X X X X Putnam 9 X X X X X X X Santa Rosa 3 X X X X X X X X Sarasota 1 X Seminole 3 X X X X X X X X Sumter 1 X X Suwanee 3 X X X X X Taylor 5 X X X X X X X X X Union 2 X X X X X X X X X Wakulla 1 X Walton 4 X X X X -59- Table 3. Black fly associations based on collections of immature stages.* T3 V n a £ ^ -* S o> k *<* <- S J5 i! a C « (B tt 03 £ S 3 tj ^ o o c o> 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. ornithophilia s. meridionale s. slossonae s. congareenarum s. dixiense s. hay si s. jenningsi s. j 'one si s. lakei s. notiale s. nyssa s. taxodiwn s. vittatwn s. decorum s. tuberosum s. vereoundum s. svp. undeterm 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 . -60- Table 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. -61- Table 4. (Continued) Date Location Collection times Species (all females) 1975: 26 Jan Double Run Creek, Site 43 1000-1330 S. s. cougar eenarvm, slossonae 12 Mar Thomas Creek, Site 159 1100-1330 s. s. congareenarum, slossonae 12 Mar 8 km west of Highway 115C on Rt 90, Site 72 1000-1400 s. slossonae 6 May Double Run Creek, Site 43 0930-1330 s. slossonae 6 May Turkey Creek, Site 216 1030-1300 s. s. slossonae, (Phosterodoros) spp. 1 June Junction SR 225/ 340, Site 8 18 June Hatchet Creek Preserve, Site 24 27-29 June 7 July 29-30 July 9-10 Aug 17-18 Sep 15-16 Oct Fisheating Creek, Site 95 Lochloosa Creek, Site 22 Fisheating Creek, Site 95 Fisheating Creek, Site 95 Fisheating Creek, Site 95 Fisheating Creek, Site 95 0930-1200 S. slossonae 1400-1930 S. slossonae, S. (Phosterodoros) spp. 0730-1145; S. slossonae, 1600-1910 S. (Phosterodoros) spp. 1730-2000 S. slossonae 0730-1200 0700-0830; 1500-1900 0830-1130; 1515-1715 0815-1315; 1620-2000 S. slossonae, S. (Phosterodoros) spp. S. slossonae, S. (Phosterodoros) spp. S. slossonae, S. (Phosterodoros) spp. S. slossonae 1976: 17 Feb Turkey Creek, Site 216 1625-1800 S. congareenarwn, S. slossonae 24 Mar Double Run Creek, Site 43 1400-1530 S. congareenarwn, S. slossonae Table 4. (Continued) -62- Date 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 -63- C. (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" 0 N D J F Months Figure 8. Seasonal occurrence of black flies in Florida. -64- 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 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 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 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 -65- CF .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) . -66- Figure 11. Lateral view of two black fly larvae (S. dixiense) Figure 12. Black fly pupa and cocoon (S. dixiense) , -67- the 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 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 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 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 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 -68- Figure 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) -69- Figure 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. -70- l\ Figure 17. The terminalia of a female S. meridionale. -71- genitalia (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 to the larvae of th e_ black flics of Florida . (Photographic 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 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 -72- 5a. (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 0 respiratory histoblast filaments with a pattern as in Fig. 53; anal tubercles small and rounded 5. jenningsi -73- 13a. (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 sub- mentum, 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 -74- A 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 -75- and 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 -76- dorsal projection (Fig. 35) S. concaveenavum 15b. Pupa with more than 12 filaments; cocoon lacking dorsal projec- tion 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, spine- like setae S. tuberosum 5b. Distimere with basal lobe bearing fine hairs only. . . S. slosscmae -77- 6a. (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 pro- jections; 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 inter- vening 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. jonesi- S. 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 -78- 10a. (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 •79- 6b. 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 -80- Introduction 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 indi- vidual 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 -81- essentially 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) in- dicated 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 sub- mental teeth (Fig. 10). The cephalic fans each contain about 60 rays with long hair-like spines. The anal tubercles are absent. -82- I Figure 18. The head spots of a larva of C. ornithophilia. Figure 19. The pupal exuviura and cocoon of C. ormithovhilia. -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 (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 -84- -• Figure 20. Tarsal claw of a female of C. ornithophilia. I Figure 21. Genital fork and terminalia of a C. ornithophilia female. -85- 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. 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 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. 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 -86- C. 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. ornithophilia- 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. 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 observations . 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 -87- identified 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. 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 j -a* Figure 22. Collection locations for C. ormithophilia in Florida. -89- Figure 23. Site 119, Gum Creek, a stream inhabited by C. ornithophilia. -90- streams 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 sub- triangular 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. ornitho- philia 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 -91- chickens. 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: -92- 18 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 (A- E.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. -93- Descrlption. 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 longi- tudinal 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 north- ern transition regions of Canada. Snow et al. (1958) captured one female. S. mcridionale at Sugar Tree, Tennessee. Stone and Snoddy (1969) -94- Figure 24. The pupa and cocoon of S. mevidionate . Figure 25. The scutum of a S. meridionale female. -95- indicate 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, pheas- ants, 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. -96- Si.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 De- Foliart (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 im- matures 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 -97- rj JJ&' Figure 26. Collection locations for S. meridionals in Florida. -98- bank 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 con- crete 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 -99- vector 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). Under- hill (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 -100- mottling and banding on the abdomen and thoracic region. The head spots are dark and distinct on a more pale head capsule. The posterior- lateral 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 cross- piece 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, mid- dorsal 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 glisten- ing 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 -101- h Figure 27. Gular notch of a 5. slossonae larva. Figure 28. The pupa and cocoon of S. slossonae. -102- m & i Figure 29. Terminalia of a male of S. slossonae. Figure 30. Terminalia of a female of S. slossonae. -103- species 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 Cycle. 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 col- lecting 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 in- habited 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. -104- Habits. 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 mid- February. 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 observat 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 -105- rJ J&** Figure 31. Collection locations for 5. slossonae in Florida. -106- of 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 ex- hibited 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 consider- able 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 -107- &r ■ '•■•■., - %4> "\v . • -'..r^ PH -tlPl WMffl Figure 32. Site 43, Double Run Creek, where S. slossonae immatures were collected. -108- eel 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 inter- mittent 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 -109- Sites 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 pre- ferences 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 in- fected with groups of tiny white spheres, probably microsporidian in- fections, on a number of occasions at just over a dozen sites. Infec- tions 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 -110- field 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 (A- D. 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), -111- 3 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 (A- J. 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), -112- 6 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: -113- 12 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 (A- G.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 (A- S.W. Frost); 1959: 10 Jan (A- S.W.Frost), 30 March (A- S.W. Frost), 7 and 18 Nov (A- S.W. Frost); 1960: 23-24 Feb (A- S.W. Frost); 1961: 6 and 17 Feb (A- S.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). -114- Lafayette 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). -115- Okaloosa 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 (A- K.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) . -116- Santa 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). -117- Wakulla 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 -118- .1 > Figure 33. Cephalic apotome of a 5. congareenavum larva. Figure 34. Venter of the larval head capsule of S. congareenamtm . -119- contains 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 res- piratory 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 sharp- pointed 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). -120- Figure 35. Pupa and cocoon of S. eongareenamm. Figure 36. Terminalia of a male of S. eongareenamm. -121- --*, , V #fr Figure 37. Terminalia of a female of 5. congareenarum. -122- Lifo 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 20- to 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). Ecology. 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. -123- Florlda observations. Stream Parameters Width Depth pH Temperature Velocity- Mean: 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 com- plete 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 -124- Figure 38. Collection locations for S. aongareenarum in Florida. -125- •* -t^ . P ft-* n, ■ ; • V .'■':: , W ■^mST'Si Figure 39. Site 216, Turkey Creek, a typical 5. aongareenarum stream. -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 col- lection. 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 occa- sions. 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, -127- and 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) . -128- Lafayette 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). -129- Taxonomy . 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). -130- Figure 40. Dorsal view of the head capsule of a S. dixiense larva. Figure 41. Gular notch and hypostomium of a S. dixiense larva. -131- The 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 iri- descent 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 orangish- red 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 -132- Figure 42. Male terminalia of S. dixiense. Figure 43. Terminalia of a female of 5. dixiense. -133- larvae and pupae into November. Garris et al. (1975) in a survey con- ducted from May through October in Sumter County, South Carolina, col- lected 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 pre- ferring 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 well- developed, large larvae and/or pupae of S. dixiense were found. The open circles designate collection sites where small larvae displaying -134- r/ J&* Figure 44. Collection locations for S. dixiense in Florida. -135- Figure 45. Site 74, Pine Barrens Creek, a stream inhabited by 5. dixiense. -136- S. 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 tem- peratures 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 -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 -133- Figure 46. Cephalic apotome of a S. haysi larva. Figure 47. Gular notch of a S. haysi larva. -139- Figure 48. Pupal exuvium and cocoon of S. haysi. -140- which 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. Distribution. 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 genera- tions 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 para- meters above summarize stream conditions prevalent when the immatures were -141- r,y JiT» Figure 49. Location of the collection site for S. haysi in Florida. -142- Figure 50. Site 195, Juniper Creek, where S. haysi was collected. -143- collected 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 sub- strate 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 num- bers 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). -144- Simulium 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). -145- =jfo«-»* " Figure 51. Cephalic apotome of a 5. jennzngsi. larva. Figure 52. Gular notch of a S. jenningsi larva. -146- Figure 53. A pupa and cocoon of S. jenningsi. Figure 54. S. jenningsi male terminalia. -147- | V ■ I Figure 55. Genitalia of a female of S. jenningsi. -148- The 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 distri- bution 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 -149- overwinter 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 men- tion 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 7.6- 9.2 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 (20- 30 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 -150- Snoddy (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 primar- ily on pupae and mature larvae with characteristic respiratory histo- blasts (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 con- stant 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 -151- r/ J0*> Figure 56. Collection locations for S. jenningsi in Florida. -152- Figure 57. Site 141 at Gulf Hammock where S. jenningsi was collected. -153- in 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 fila- ment 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 dif- ference 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 -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). -155- Putnam 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 ex- uvium and cocoon has been deposited in the U.S. National Museum. Description. 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 dis- tally and nine additional filaments which arise from the basal one (Fig. 60). -156- ft Figure 58. Cephalic apotome of a S. jonesi larva. Figure 59. Gular notch of a S. jonssi larva. -157- Figure 60. Respiratory organ of a S. jonesi pupa. -158- The male is black and velvety in appearance with a pair of triangu- lar, 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 collec- tions 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 history. 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. Habits. 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. -159- Figure 61. Terminalia of a male of S. jonesi. Figure 62. Genital fork and terminalia of a female of 5. jonesi. -160- Florida 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, -161- rJ *P» Figure 63. Florida collection locations for 5. jonesi. -162- Figure 64. Site 210, the Fenholloway River, a S. jonesi collection location. -163- sedges, 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 popula- tions 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 associ- ations. 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 •164- S. 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 Tor- reya 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 fila- ments 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. -165- Petioles 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 -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). -167- 57) 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). -168- Liberty 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); -169- 1975: 28 March (S), 12 June (S,L). Taylor Co. 210) 1954: 8 April (Larvae, Pupae- CM. 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 yellow- brown head capsule. The pattern of the spots on the cephalic apotcme -170- is 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, -171- Figure 65. Cephalic apotome of a S. lakei larva. ■Mr Figure 66. Gular notch and hypostomium of a 5. lakei larva. -172- Figure 67. Pupa and cocoon of S. lakei. Figure 68. Dorsal view of a male of S. lakei. -173- Figure 69. Terminalia of a male of S. lakei. Figure 70. Terminalia of a female of S. lakei. -174- pupae 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 per- manent 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 characteris- tically near neutrality or just below in pH reaction and all are -175- Figure 71. Collection locations for 5. lakei in Florida. -176- Figure 72. Site 135, Otter Creek, a collection site for S. lakei. -177- moderately 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. Im- matures 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 popu- lations 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 -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 labora- tory. 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 else- where 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 respira- tory 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 ter- minalia 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 -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). -180- Hardee 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 -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 -182- to 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 hori- zontally 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 three- fourths 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. -183- Figure 73. Head spots of a S. notiale larva. Figure 74. Gular notch of a S. notiale larva. -184- Figure 75. Pupal exuvium and cocoon of S. notiale. Figure 76. Scutum of a male of 5. notiale. -185- / i Figure 77. Terminalia of a male of 5. notiale. Figure 78. Terminalia of a female of S. notiale. -186- Distribution. 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 medium- sized streams with high quality water exhibiting a temperature of 13- 16°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) -187- r/ J*** Figure- 79. Collection locations for S. notiale in Florida, -188- Figure 80. Site 88 at Chattahoochee where S. notiale immatures were found. -189- high. 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 popula- tions of S. notiale were found attached to outcrops of the yellow rock- like 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, -190- Lee 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 examina- tion 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. -191- Figure 81. S. nyssa pupa and cocoon. -192- Life 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 col- lected on river weed, Podostemon ceratophyllum, in shallow rapids (.76- 1.06 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 -193- Figure 82. Collection locations for 5. nyssa in Florida. -194- Figure 83. Site 116, Blue Creek, where S. nyssa was collected. -195- S. 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 pre- sented 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. nyssa. Holmes Co. 116) 1975: 19 Jan (P) . Orange Co. 171) 1947: 9 Jan (P - H.K. Gouck). Simu Hum (Pkoeterodor-os) taxodium Snoddy 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 -196- associated 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. Para- types 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 later- ally and sparce, small golden hairs. The subcosta is bare. The cerci are broadly rounded distally, bear many setae and in lateral view appear -197- Figure 84. Cephalic apotome and head spots of a S. taxodium larva. Figure 85. Gular notch of a S. taxodiiun larva. -198- Figure 86. Pupal exuvium and cocoon of S. taxodium. % Figure 87. Male terminalia of S. taxodium. -199- > Figure 88. Female terminalia of S. taxodium. -200- as 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 neu- tral (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 -201- Figure 89. Collection locations for S. taxodium in Florida. -202- October 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 -203- Figure 90. The Ichetucknee River, a S. taxodiwn collection site. -204- usually 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). -205- Columbia 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) . -206- Levy 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) . -207- Simuliwn 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 speci- men at the University of Lund and designated it as the lectotype in the event other of Zetterstedt ' s specimens materialize. Lugger (1897) de- scribed a male, larva and pupa for this species under the name S. tribu- latum. 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 -208- developed. 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 es- pecially prominent medial and lateral teeth and the margins of the sub- mentum beyond the lateral teeth are conspicuously serrate. There are four to five long and additional short setae on each side of the sub- mentum. The antenna is rather short with two white spots. In alcohol the abdomen is gray or greenish-gray to gray black with lighter inter- segmental 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 -209- Figure 91. Head spots of a S. vittatum larva. Figure 92. Gular notch of a S. vittatum larva. -210- Figure 93. S. vittatum pupa and cocoon. {?** Figure 94. Terniinalia of a 5. vittatum male. -211- Figure 95. Scutum of a female of 5. vittatum. < ..'• >*A Figure 96. Terminalia of a female of S. vittatum. -212- in 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 pro- minent 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, re- lease eggs, and then fly up and repeat the process. Peterson (1961) -213- found 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 sub- strate 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 desic- cation (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), -214- four 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 abun- dant 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 silt- covered 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. -215- Jamnback (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 -216- dependent 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 situ- ation 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 Withla- coochee River, Madison Co., which I examined but did not collect. Simulium vittatum is the seventh most widespread black fly in the state -217- r/ J0 Figure 97. Collection locations for S. vittatum in Florida. •218- occurring in the wesLern, central and more southern sections. There are five types of flowing water in which the immatures occur: 1) out- flows 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 exceed- ing 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 tem- perature 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 in- crease the amount of abrasive particulate matter present possibly detri- mental 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 -219- Figure 98. Site 167 at Crestview where S. vitiation was collected. -220- eggs 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 loca- tion, 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 con- stricted 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 -221- found 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 char- acteristics 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 •222- with 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 alco- hol, 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 -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). -224- Nassau 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). -225- Taxonomy. 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 un- clear. Stone and Snoddy (1969) suggest S. argyreatum Meigen, with which 5. noelleri- Frederichs 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 -226- Figure 99. Head spots of a S. decorum larva. ■*+ Figure 100. Gular notch of a 5. deaonan larva. -227- Figure 101. S. decorum pupa and cocoon. Figure 102. Male terminalia of S. decorum. -228- Figure 103. Female terminalia of S. decorum. -229- scape 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, sub- apical 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 de- velop 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 genera- tions in South Carolina from February to May and suggest that with suf- ficient rain S. decorum breeds all summer with a generation every month. Stone and Snoddy (1969) indicate the initial mass emergence of adults -230- occurs 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 tem- peratures 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 pre- ferentially 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) in- fections in S. decorum. McKague and Wood (1974) found a granular SRIO Altosid formulation at .1 ppm concentration was 100% effective in halt- ing 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 pro- tected 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 -231- Peterson, 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 develop- ing 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, -232- rr _«^ ^fit. -^ Figure 104. Collection locations for 5. decorum in Florida, -233- respectively. 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 popu- lations of larvae and pupae were found, especially during March, concen- trated 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 tap- ping 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 -234- Figure 105. Shepard's Mill, Site 86, where S. decorum was collected, -235- Crestview 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). -236- Simulium 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. 0 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 in- distinct 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 sub- esophageal ganglion is dark. The anal tubercles are small and incon- spicuous. -237- Figure 106. The cephalic apotome of a S. tuberosum larva. Figure 107. The gular notch of a S. tuberosum larva. -238- Figure 108. Pupa and cocoon of S. tuberosum. * Figure 109. Male terminalia of S. tuberosum. -239- v?aft Figure 110. Terminalia of a female of S. tuberosum. -240- The pupa is 2.5-3 mm long in a fairly uniformly textured, light- brown, 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 -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 pre- sent 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 occur- ring 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 -242- females 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 ParametersJ 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. -243- Figure 111. Collection locations for S. tuberosum in Florid; -244- of 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. -245- The 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 num- bers 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 con- crete, 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-bear- ing 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 -246- Figure 112. Site 56 in Clay County, a collect! on spot for S. tuberosum. -247- numbers 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 col- lected 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 -248- the 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 collection 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 (A- G.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 (A- J. 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: -249- 2 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 jan (S,M,L,P5C,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 -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 (A- J. 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) -251- Clay 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: -252- 18 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) 1975: 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) i974: 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,M5L, 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: -253- 17 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) . -254- Okaloosa 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). -255- Seminole 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) -256- indicates S. verecundum may be the same as S. argyreatum Meigen as de- fined 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 an- teriorly (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 well- woven . 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 -257- Figure 113. The head spots of a S. verecundwn larva. Figure 114. The gular notch of a S. verecundwn \t -258- Figure 115. The pupa and cocoon of S. vevecundvm. Figure 116. Male terminal ia of S. vereaundum. -259- Figure 117. Terminalia of a female of S. veveoundum. -260- from 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. vere- aundum eggs overwinter. Larvae appear in late April or early May and adults are present by late May. Anderson and Dicke (1960) found imma- ture 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% -261- parasitism 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 -262- -/ .^* Figure 118. Collection locations for S. verecundum in Florida -263- records 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 up- stream. 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 -264- Figure 119. Panther Creek, Site 223, where S. verecundum was collected. -265- range of plants in the streams in addition to twigs and, on a few occasions, to the rock-like clay substrate of a stream. When construc- tion at Sites 6 and 17 destroyed sections of the streams rich in vege- tation 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, tempera- ture 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 col- lected 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. -266- Florida 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). -267- Hamilton 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) . -268- Cnephia 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 elong- ate, 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 num- ber 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. -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. -270- Figure 122. Collection location for Cnevhia species No. 1 in Florida. -271- Simuliion 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, well- woven, 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 pre- sent 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 ex- tends 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. -272- The 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. -273- Figure 123. Pupal exuvium and cocoon of Simuliwn species No. 1, -274- Figure 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 -275- rJ J&* Figure 126. Collection location for Simuliwn species No. 1 in Florida, -276- Figure 127. Small flow to Pine Barrens Creek, Site 74, where Simulium species No. 1 was collected. 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 semi- wild 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 in- fected 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- -278- Table 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 Junc- tion (Site 8) 9-11 May 1975 negative 9163, 9167, cage 9170 Hatchet Creek Preserve (Site 24) 17-23 May 1975 only 9170 posi- t ive , 2 June 245, 246, cage 247 SR 225/340 Junc- tion 16-22 June 1975 only 245 posi- tive, 11 July 223, 224, cage 312 Lochloosa Creek 2-10 July 1975 only 223 posi- tive, 20 July 313, 314, ramp trap 315 Hatchet Creek Preserve 3-10 July 1975 only 315 posi- tive, 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 Junc- tion 4-8 March 1976 (birds stolen) 483, 484, cage 485 SR 225/340 Junc- tion 9-16 March 1976 483, 484, 485 all positive, 22 March -279- birds (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 in- fections 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 -280- placed in similar cages at ground level remained negative for the dis- Results 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 Pkoatero- dovos 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 58- 90% 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 un- covered condition. The ramp trap was hoped to be an improvement •281- Table 6 . Black flies captured in ramp traps. Turkey Number Location Dates Captures (all 1975) (all females) 9170 Rt 225/340 June- 21-22 June tion 149, 150, Lochloosa Creek 29-30 April 0 175 (Site 22) 9172, 9168, Rt 225/340 June- 9-11 May 0 9169 tion (Site 8) 9170 ' Hatchet Creek 18-20 June 0 Preserve (Site 24) 1 S. slosi 313, 314, Hatchet Creek 3-10 July 315 Preserve 2 S. slossonae -282- Table 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 0 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 0 6 July 0830-0940 S. slossonae 18 Sept 0900-1.130 0 -283- Table 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, 9966 Lochloosa Creek 320, 9966 SR 225/340 Junction (Site 8) 324 Double Run Creek 324 Turkey Creek (Site 216) 479, . 480, Lochloosa Creek 481 476, • 477, SR 225/340 Junction 478 485 Double Run Creek 485 Turkey Creek 483 Sante Fe College (Site 28) 21 April 1975 1600-1700 0 22 April 1200-1500 0 5 July 1700-1800 S. slossonae 6 July 0945-1110 S. slossonae 19 July 0830-1100 S. slossonae 22 July 0830-1030 S. slossonae 29 July 0930-1100 0 5 Aug 1030-1225 S. slossonae 30 Aug 1000-1200 0 30 Aug 1740-1930 S. slossonae 7 Jan 1976 1330-1510 S. slossonae 7 Jan 1600-1715 S. eongarecnarwn, S. slossonae 4 March 1600-1630 S. conaareenarum, S. slossonae 4 March 1500-1530 S. congareenamen, S. slossonae 24 March 1400-1600 S. slossonae 24 March 1645-1830 0 26 March 0930-1030 0 485 SR 225/340 Junction 31 March 1600-1730 S. slossonae -284- in that it still used turkeys as attractants; however, flies entering to feed were considered captured and the col lector could recover speci- mens 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 orni- thophilic 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 de- veloped 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 ex- periment 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, cling- ing 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. -285- Perhaps 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. aongaveen- avum, 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 thun- derstorms 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 -286- numbers 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. mevi- dionale adults were reared. In addition, reared females of the S. (Phos- tevodovos) species jenningsi, jonesi, and notiale and undetermined wild- caught 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 £UeS 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 -287- they 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 tem- peratures (13 -32°C - 55 - 90°F) and relative humitidies (40-90%). Un- interrupted feeding times for S. slossonae ranged from 1 to 10 minutes with an average of 4.3 minutes. Simuliwn congareenar-um fed at tempera- tures 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 pro- ceeded to engorge in three minutes (1729-1732 hrs) . Cnephia ovnitho- philia, 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 -288- perpendicular 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 ex- tensively than any other species with the abdomen expanding to about three tiraes 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 leuco- cytozoonosis 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 innocu- lated 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 <-t 00 60 60 Cu -289- co cm m co oo co oo co a. in cnj en cm en vD in co a> r^ en cm CM CM CM eneneoo->CMcn a\ cm vo m in m H O^ U-> CT\ to 60 oo a. ex ex, \D Csl r-{ \£> r*- r^ Ln in o ^o ^o \o cm en ,-i tn i£5 vD VD rH cm en VD CM CM CM CM »cf ct\ cn cn cn cn m m in m m in 00 00 CO 00 CO T3 T3 T3 ~0 WQ>Q>IUQ)Q) o o a o o o ^ S s o o o o o o CO CO. <0 Co Cq CQ Co tQ Co Co Co CO Cq tQ Co Co Co Co Co O r-t cm en -290- S. jonesi, and three C. omithophilia fed twice but no L, smithi infec- tions 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 (March- April). 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 inter- rupted 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 trans- mission 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 -291- Figure 128. Gametocytes (G) of L. smithi among normal turkey blood cells. Figure 129. Ookinetes of L. smithi. -292- 20u Figure 130. Sporozoites of L. smithi photographed in sal: figure 131. Stained sporozoites of L. smithi. -293- elongate sporozoites packed like toothpicks as illustrated for other species of Leucooytozoon (Pallia and Bennett, 1961a) were never posi- tively 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 ex- cept 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. 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 identi- fied 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, espe- cially 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- -295- as vectors of L. smithi to turkeys in Florida. In order of de- creasing importance, these species are: S. slossonae , S. oongare- enarum, 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 loca- tions miles away from black fly streams or force screened outdoor or indoor rearing procedures. 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. 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