THIE BAICK FLIES (DIPTERA:: SIHULLIIDAE) OF FLOIDAT, AN1\D 'THEIR
INVOLVEMENT INi THE TRANSMISSION OF'
L~eucocytozoon smith TO TURKEYS
DENNIS DREW PINKOVSKY
A D)ISSERTATIOJN PRESENTED TO THE GRA.DUAT~ COUNCIL OF
THE UNIVERSrTY OF FLOREDAl
IN PARTIAL FULFILLMENTT UF THE REQUIREMENTS FOR\ TbE
DEGREE OF DOCTOR OF PHILOSOPHti
UNIZVERSITY OF FLO;:rIDA
I sincerely thank Dr. J.F. Butler, my Committee Chairman, for his
constructive suggestions and support during my research endeavors and
for his valuable, critical review of my dissertation.
To Dr. D.F. Forrester I extend my deep appreciation for the advice
and guidance he enthusiastically offered during my transmission studies
and for the equipment and facilities he generously allowed me to use.
To all the mem~btrs of my Ph.D. Committee I express appreciation
for their critical appraisal of my dissertation and their helpful
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
I express my gratitude for the opportunity to examine black fly
specimens collected in Florida which were made available to me by
numerous individuals and institutions.
To the staff at the U.S. Department of Agriculture Insects Affecting
Man Laboratory, Gainesville, I extend my appreciation for the use of equip-
ment and for advice during my research and academic studies.
I am grateful to Dr. E.V. Komarek and the staff at the Tall Timbers
Research Station and L..E. pjilliams, D.H. Austin, and T. Peoples of the
Florida Game and! Fresh Water Fish~ Commission for the use of facilities
and other assistance during my collecting trips.
I wish to also thank the State of Florida Division of Recreation
and Parks for collecting permits and the opportunity to gather specimens
at the beautiful State Parks around Florida.
I extend my deep gratitude to the U.S. Air Force Institute of
Technology Civilian Institutions Program for financial support and for
the opportunity to attend the University of Florida and pursue my Ph.D.
To P. Humphrey, D. Young, L. DuBose and especially T. DiNuzzo I
express my sincere thanks for good-spirited support during my research.
General Comments .
Introduction to the Black Fly Keys. ..... .
A key to the larvae of the black flies of
Florida . . . . . *
A key to the pupae of the black flies of
Florida . . . . . . .
TABLE OF CONTENTS
LIST OF TABLES. ...c.
LIST OF FIGURES .....
AB STRACT. ........
I. INTRODUCTION. ..........
II. LITERATURE REVIEW ........
Taxonomy and Distribution. .
Damage .. . .. . .. .
Florida Ecological Habitats ...
Leucocytozoon smith. ......
III. MATERIALS AND METHODS ,.....
Black Fly Survey. ........
Leucocy-tozoon smithi Transmission
IV. RESULTS AND DISCUSSION. .....
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TABLE OF CONTENTS
A key to the adult male black flies of
Florida . . . . . . . .
A key to the adult female black flies of
Florida . . . . . . . . .
Introduction to the Individual Species Sections...
Chephia (Cnephia) ornithophilia Davies, Peterson,
and Wood . . . . . . . .
Simutiumr (Byssodon) meridionaie Riley. ......
Simulium (Bt~ssodon) slo~ssonae D~yar and Shannon ..
Sirmuium (Eurshnutium) cong~areenatum(Dyar and
Shannon) . . . . . . . . .
Simutium (Phosterodoros) dixieinse Stone and Snoddy
Simutium (Phosterodoros) haysi Stone and Snoddy..
Simu~ium (Phosterodoros) denningsi Malloch.. ..
Simutlium (Phosterodoros) jonesi Stone and Snoddy..
SimuZium (Phosterodoros) lake~i Snoddy.. .....
SimuZium (Phoster~odoros) notiale Stone and Snoddy.
SimuZium (Phosterodoros) nyssa Stone and Snoddy..
SimuZium (Phosterodoros) taxodium Snoddy and
Beshea-r. . . . . . . . .
Simu*Zium (Psilozia) vistatum Ze~ttrstedt
Simulium (Simutium) decorun Walker ........
Simulium (SimuZium) tuberosum (Lundstram)......
SimuZium (S~imulium) verecundwnr Stone and Jamnback..
Opephia species Undetermined No. 1.. ......
Shaulium species Undetermined No. 1.... ....
Leucocytozoon smith Transmission... ........
V. CONCLUSIONS.. ... .. .......
LITERATURE CITED. ...... ........
APPENDIX. COLLECTION SITE NAMES AND LOCATIONS........ .
BIOGRAPHICAL SKETCH.. ... .............
LIST OF TABLES
1. Florida black fly species. .. .. .. .. .. ... 55
2. Florida black fly distribution records by county .. .. 57
3. Black fly associations based on! collections of immature
stages .. .. .. ... .. ... .. .. ... 59
4. Black flies captured in Manitoba traps . ... .. .. 60
5. Sentinel turkey locations and results. .. .. .. .. 278
6. Black flies captured in ramp traps .. .. .. .. .. 281
7. Blackout box trapping results. . .. ... .. ... 282
8. Black fly captures from exposed turkeys. .. .. .. .. 283
9. Leucocytozoon smith transmissions .. .. . ... 289
LIST OF FIGURES
1. A modified Manitoba trap with a black plastic skirt . .. 44
2. Sentinel turkeys in an exposure cage. .. .. .. .. .. 47
3. A blackout box trap in the field. .. . .. ... ... 47
4. One view of a ramp trap .. .. .. ... ... .. 48
5. An exposed turkey in the field. .. .. ... .. .. 48
6. Glass container and paper cartons used for holding
black fly adults alive in the laboratory. .. .. .. .. 51
7. Locations in Florida where black flies have been
collected . .. .. . .. . .. .. .. 56
8. Seasonal occurrence of black flies in Florida .. ... 63
9. Dorsum of the head capsule of a black fly larva
(S. slossonae) .. .. .... .. .. .. .. .. 65
10. Venter of the head capsule of a black fly larva
(C.ornithoph~iia ... .. .. . .. .... .. .. 65
11. Lateral view of two black fly larvae (S. dirEense) .. .. 66
12. Black fly pupa and cocoon (S. diriense). .. .. . ... 66
13. A wing of the black fly onephia ornith~ophilia .. .. .. 68
14. A frontal view of the head of a female black fly
(S. notiate). .. .. .... .. .. . ... . .. 68
15. The male genitalia of a black fly, Cnephia
ornithoohilia ... .. .. .. . ... ... . .. 69
16. The distal portion of the hind leg of a S. meridionate
female. .. ... .. ... .. .. .. . .... 69
17. The terminalia of a female S. merZidonatea .. .. .. .. 70
18. The head spots of a larva of C. ornithophiflia . ... .. 82
LIST OF FIGURES
19. The pupal exuvium and cocoon of C. ornithophilia. . ... 82
20. Tarsal claw of a female of C. ornithophilia .. .. .. 84
21. Genital fork and terminalia of a C. ornithophilia female. 84
22. Collection locations for C. ornithophilia in Florida. .. 88
23. Site 119, Guim Creek, a stream inhabited by
C. ornith~ophilia. .. . ... .. . . .89
24. The pupa and cocoon of S. mer~idionate .. ... . 94
25. The scutum of' a S. meridionale female . .. .. 94
26. Collection locations for S. meridionale in Florida. .. 97
27. Gular notch of a S. sZossoncae larva .. .. .. .. .. 101
28. The pupa and cocoon of S. slossonae . ... .. .. .. 101
29. Terminalia of a male of S. s~ossonae. .. .. .. .. .. 102
30. Terminalia of a female of S. sZossonae. .. .. .. .. 102
31. Collection locations for S. slossonzae in Florida. .. .. 105
3.Site 43, Double Run Creek, where S. so~;ssonae
immatures were collected. . .. .. .. .. .. 107
33. Cephalic apotome of a S. congareenarum larva. . .. ... 118
34. Venter of the larval head capsule of S. congareenaumo .. 118
35. Pupa and cocoon of S. congareenarum .. .. ... .. 120
36. Terminalia of a male of S. congareenarum. .. .. ... 120
37. Terminalia of a female of S. congaree~nawn. .. .. .. 121
38. Collection locations for S. congereencrum in Florida. .. 124
39. Site 216, Turkey Creek, a typical S. congareenarum
stream. .. .. ... .. .. .. .. .. .. ,.. 125
40. Dorsal view of the head capsule of a S. didcense larva. . 130
41. Gular notch and by~postomiumn of a S. d'ixiense larva. . .. 130
LIST OF FIGURES
Male terminalia of S. diziense. ..........
Terminalia of a female of S. dizciense .......
Collection locations for S. dizciense in Florida ..
Site 74, Pine Barrens Creek, a stream inhabited by
S. didcense . . . . . . . . . .
Cephalic apotome of a S. haysi larva. .......
Gular notch of a S. haysi larva ..........
Pupal exuvium and cocoon of S. haysi. .......
Location of the collection site for S. hacysi in Flol
Site 195, Juniper Creek, where S. haysi was collect~
51. Cephalic apotome of a S. jenningsi larva. .. .. ... 145
52. Gular notch of a S. jenningsi larva . ... .. .. .. 145
53. A pupa and cocoon of S. jenningsi .. .. .. .. .. .. 146
54. S. jenningsi male terminalia. .. ... .. .. .. .. 146
55. Genitalia of a female of S. jenningsi .. .. .. . 147
56. Collection locations for S. jenn~ingsi in Florida. . ... 151
57. Site 141 at Gulf Hamrmock where S. jennings~i was collected 152
58. Cephalic apotome of a S. jonesi larva ... .. .. 156
59. Gular notch of a S. jonesi larva. .. .. .. . .. 156
60. Respiratory organ of a S. jonesi pupa .. .. .. . 157
61. Terminalia of a male of S. jonesi .... .. .. .. 159
62. Genital fork and terminalia of a female of S. jon~esi. .. 159
63. Florida collection locations for S. jonesi. .. .. .. 161
64. Site 210, the Senholloway River, a S. jcnesi collection
location. ... .. .. .. .. .. .. .. .. .. 162
LIST OF FIGURES
Cephalic apotome of a S. lakei larva. ........
Gular notch and hypostomium of a S. lakei larva ...
Pupa and cocoon of S. lakei .............
Dorsal view of a male of S. laket .....
Terminalia of a male of S. lakei....... .. .
Terminalia of a female of S. lakei. .........
Collection locations for S. lakei in Florida. ....
Site 135, Otter Creek, a collection site for S. lakei
Head spots of a S. notiale larva. ..........
Gular notch of a S. notiate larva ..........
Pupal exuvium and cocoon of S. notiale. .......
Scutum of a male of S. notiate. ...........
Terminalia of a male of S. notiale. .........
Terminalia of a female of S. notiaie. ........
Collection locations for S. notCate in Florida. ...
80. Site 88 at Chattahoochee where S. notiate immatures
were found. .. ... .. .. .. . .. .. . 188
81. S. nyssa pupa and cocoon. .. ... .. ... .. .. 191
82. Collection locations for S. nyssa in Florida. ... .. 193
83. Site 116, Blue Creek, where S. nyssa was collected. . .. 194
84. Cephalic apotome and head spots of a S. taxcodium larva. . 197
85. Gular notch of a S. tarcodium: larva. .. ... .. .. 197
86. Pupal exuvium and cocoon of S. taxcodiu7 . ... .. .. 198
87. Male terminalia of S. txodium. . .. .. .. ... .. 198
88. Female terminalia of S'. Laod~ium. .. .. .. .. .. .. 199
110. Terminalia of a female of S. tCuberosumr. .. ... . .. 239
111. Collection locations for S. tubrerosum in Florida. .. .. 243
112. Site 56 in Clay Countyr, a collection spot for S. tuberosum. 266
LIST OF FIGURtES
89. Collection locations for S. tax~odium in Florida
90. The Ichetucknee River, a S. taxcodium collection
91. Head spots of a S. vittatum larva .......
Gular notch of a S. vittatum larva. ............
S. vittatum pupa and cocoon ................
Terminalia of a S. v-ittatum male. .............
Scutum of a female of S. vittatum.... .....
Terminalia of a female of S. vittatum... ....
Collection locations for S. vittatum1 in Florida ......
Site 167 at Crestview where S. vittatum was collected ...
Head spots of a S. decorum larva. .............
Gular notch of a S. deconrum larva .............
S. decorum pupa and cacoon. ................
Male terminalia of S. decoum ..... .....
Female terminalia of S. decorum ..............
Collection locations for S. decorum in Florida. ......
Shepard's Mill, Site 86, where S. decorum was collected ..
The cephalic apotome of a S. tuberosum larva. .......
The gular notch of a S. tuberosum larva ..........
Pupa and cocoon of S. tuberosum.... .....
Male terminalia of S. tuberosum.... .....
LIST OF FIGURES
113. The head spots of a S. verecundum larva. .. .. .. .. .257
114. The gular notch of a S. vereoundum larva .. . .. ... .257
115. The pupa and cocoon of S. verecundum .. . .. .. .258
116. Male terminalia of S. verecundum .. .. .. .. .258
117. Terminalia of a female of S. vereoundum. .. .. .. .. .259
118. Collection locations for S. verecundum in Florida. . .. .262
119. Panther Creek, Site 223, where S. verecundumlu was collected .264
120. Cephalic apotomce of a larva of Onephia species No. 1 .. .269
121. Venter of the head capsule of a larva of Cnephia species
No. 1... . .. .. .. .. . . .. .. .. 269
122. Collection location for Q~nephia species No. 1 in Florida .270
123. Pupal exuviumn and cocoon of SimuZi~um species No. 1 .. .273
124. Larval head spots of Simulium species No. 1. .. .. .. .274
125. Venter of the head capsule of the larva of SimuZium species
No. 1. .. .. .. .. .. .. .. ... .. ... .274
126. Collection location for SimuL~ium species No. 1 in Florida. .275
127. Small flow to Pine Barrens Creek, Site 74, where
Simulium species No. I was collected .. .. . .276
128. Gametocytes (G) of L. smith among normal turkey blood
cells. .. .. .. .. . . . .. .. .291
129. 00kinetes of L. smith .. .. .. . . .291
130. Sporozoites of L.. smith photographed in saline. . ... .292
131. Stained sporozoites of L. smithi . . .. .. . .292
Abstract of Dissertation Presented to the Graduate Council
of the University of Florida in Partial Fulfillment of the Requirements
for the Degree of Doctor of Philosophy
THE BLACK FLIES (DIPTERA: SIMULIIDAE) OF F~LORIDA AZND THEIR
INVOLVEMENT IN THIE TRANSMISSION OF
Leucocytozoon smith TO TURKEYS
Dennis Drew Pinkovsky
Chairman: Jerry F. Butler
Major Department: Entomology and Nematology
Immature and adult black flies (Diptera: Simuliidae) were collected
in Florida over a period of three years. Eighteen species of black
flies including ten which are new records for the State were found to
occur in Florida. Records of black flies from 192 locations in 50
counties are included. Biological information is provided for each
species together with data on distribution, seasonal occurrence, scream
ecology and species associations. Keys to the Simuliidae of Florida are
provided and structures are illustrated. Representative specimens have
been deposited in the Florida State Collection of Arthropods and in the
United States National Miuseum, Washington D.C.
Transmission investigations have incriminated three species,
SimuZiumi congareenrum~, S. me~ridion~ale, and S. slossonae, as vectors of
Leucoceytozon smithi to turkeys in Florida. On nineteen occasions
L. smithli was transmitted to domestic turkeys by the bites of infected
Members of the family Simuliidae as immatures inhabit flowing water
and as adults are often blood feeders which may vector diseases to man
and animals in many parts of the world. A knowledge of the composition,
distribution, ecology, and habits of the simuliid fauna of any area of
local concern is essential for proper assessment of the impact on man of
these insects. Successful black fly control is dependent on thorough
knowledge of the major breeding sites, seasonal occurrence and other
facts concerning local Simuliidae.
When I arrived in Florida to begin my Ph.D. research I wrote to
the Florida Division of Health offices in Jacksonville, Vero Beach, and
Panama City and enquired about previous investigations on the black
flies of Florida. I found that little work had been done in the State
on this family of biting flies (Beck, 1973; Linley, 1973; Rogers, 1973 -
all personal communications). Mrs. A.T. Slosson collected black flies
in Florida (Dyar and Shannon, 1927), probably at the turn of the
twentieth century. Some of Mrs. Slosson's specimens and a portion of
those of Calvin Jones, Harry Couck, Darrell Anthony and other USDA
researchers who collected black flies in the northern and central por-
tions of the State in the 1940's and 1950's are located in the U.S.
National Museum. Based at least in part on the examination of
these specimens Stone (.1965.) listed the following seven species from
Florida: Crnephia (Cnephia) pecuarum (Riley); Simulium (Eusimuli~un)
congareenatnum(Dyar and Shannon); Simulium (Byssodon) meridionaite Riley;
Simuliwn, (Byssodon) saossonae Dyar and Shannon; Simulium (Simutium)
decorum Walker; SimuZitrm (Simulium) jenningsi Malloch; and Sirmutium
(Simutium) tuberosum (Lundstrijm). Stone and Snoddy (1969) did not list
S. jenningsi as occurring in Florida but did record two new species
Simulium (Phosterodoros) jonesi Stone and Snaddy and Simulium (Phostero-
d'oros) nyssa Stone and Snoddy from the State. Thus the total number
of species of black flies known to occur in Florida when I began my
research was eight.
This research was initiated in the fall of 1973 with the following
objectives: 1) to determine the species complement, distribution and
seasonal occurrence of black flies throughout Florida; 2) to gather
ecological information primarily on the immature stages; 3) to provide
keys with illustrations to the black flies of Florida; 4) to gather and
deposit in the U.S. National Museum of Natural History and the Florida
State Collection of Arthropods as complete a collection of adult and
immature Florida black flies as possible; and 5) to determine the vec-
tors of Leucocytozoon smith in turkeys in Florida.
Taxonomy and Distribution
Rubtsov (1974) presents a concise discussion on the history and
major advances in black fly taxonomy. Linnaeus (1758) first described
two simuliids and assigned the names CuZez reptan~s and Culex: equinzus
but did not differentiate them from mosquitoes (Davies et al., 1962).
Latreille created the name Simulium for the genus in 1802 using Rhagio
cotomb~aschenzsis Fabricus as the type (Stone and Jamnback, 1955). This
remained the only genus for the thirty to forty black fly species de-
scribed up to the turn of the twentieth century. The work of Smith and
Kilbourne (1893) and others which focused on arthropods as vectors of
diseases stimulated much research on insects and related groups of
medical and veterinary importance. Roubaud (1906) designated two black
fly subgenera, Prosimuium and Eusinulium, based on differences in wing
venation. Lundstram (1911) in Sweden and Jobbins-Pomeroy (1916) in the
United States were early taxonomists who made use of the male terminalia
of simuliids to separate species. Edwards (1915) in England also used
male terminalia but more significantly in 1920 stressed the importance
of studying black fly larvae and pupae. Dyar and Shannon (1927) first
used female genitalia to separate North American black flies. Enderlein
(1930) divided the family Simuliidae into 29 genera. Smart (1945) took
a more conservative approach and distinguished 6 genera in the family:
ParasimuZium Malloch, ProsirmuLium Roubaud, Aulstrosimulium Tonnoir,
Gigantodcw: Enderlein, Onephia Edwards and Simulium Latreille. Stone
(1963) listed 11 genera and 22 subgenera in the Simuliidae which he
considered valid. Crosskey (1969) stressed a trinomial approach to
simuliid taxonomy with heavy reliance on subgenera. Rubtsov (1974)
recognized 17 genera of black flies in the Palaearctic region and listed
59 genera for the Simuliidae of the world, many of which American and
British authors regard as subgenera.
Brues et al. (1954) provide a worldwide bibliography of important
papers which deal with the taxonomy of the Simuliidae up to the early
1950's. In addition to the works already mentioned outstanding publi-
cations since about 1950 dealing with the classification and distribu-
tion of black flies outside of the United States include:
Palaearctic region Davies (1968) Britain, Carlsson (1969) Spain,
Rubtsov (1956 and 1962) U.S.S.R., Kuusela (1971) Finland,
Zivkovic (1971) Yugoslavia, Rivosecchi (1971 and 1972) Italy,
Crosskey and Pete~rson (1972), and Zwick (1974) Germany;
Ethiopian region Crosskey (1957) West Africa, Travis et al. (1967),
Crosskey (1969) Africa and its islands, Fain and Elsen (1973) -
Cameroons, Lewis and Raybould (1974) Tanzania;
Oriental region Travis and Labadan (1967a) Asia and European U.S.S.R.,
Delfinado (1969, 1971) Philippines, Crosskey (1973), Takaoka
(1973) Nansei Is., Uemoto et al. (1973) Japan, Lewis (1973) -
Pakistan, Datta (1973, 1974, 1975) India;
Australian Dumbleton (1963, 1972) Australia and New Zealand, Travis
et al. (1968);
Neotropical Leon and Wygodzinsky (1953) Ecuador, Dalmat (1955) -
Guatemala, Vulcano (1967), Travis and Labadan (1967b),Barreto
(1969) Colombia, Wygodzinsky and Coscaron (1970, 1973), Wygod-
zinsky and Najera (1970), Wygodzinsky (1971) Northern Andes,
Perez (1971) Venezuela, Rubtsov and Avila (1972) Cuba, Travis
et al. (1974);
Nearctic (excluding the U.S.) -- Syme and Davies (1958), Davies et al.
(1962) Ontario, Wood et al. (1963) Ontario, Travis et al. (1969),
Peterson (1970) Canada and Alaska, Lewis and Bennett (1973) -
In addition to the above works those of Rubtsov (1970, 1974) and
Coscaron and Wygodzinsky (1973) mention the variability which has been
observed in characters that are used for taxonomic determination of
black flies. Gambarian and Terterian (1973) applied a numerical tax-
onomy approach and using 100 characters tried to separate simuliids in
the Eusimulium group. The subgenus was found to be very homogeneous
and creation of supraspecific taxa was not statistically justified.
Internationally, black fly species determinations based on classical
external morphological characters and behavioral differences are being
supported by cytological, specifically chromosomal, studies. Rothfels
(1956) provided an introduction and overview on analytical techniques
for cytologically comparing black flies. Landau (1962), Dunbar (1969),
and Vajime and Dunbar (1975) used chromosomal differences to separate
forms or sibling species of a variety of simuliids including disease
vectors where previously each had been considered a single species.
Taxonomic works covering the whole United States include: Coquillett
(1898, 1902), Malloch (1914), Jobbins-Pomeroy (1916), Dyar and Shannon
(19217), and Stone (1965). Other publications were more regional in
scope. Western and midwestern articles include: RTinn (1938), Smith
and Lowe (1948) California; Nicholson and M~ickel (1950) Minnesota;
Stone (1952), Somrmerman (1953) Alaska; Wirth and Stone (1956) -
California; Stone and DeFoliart (1959); Anderson and Dicke (1960) -
Wisconsin; Stone and Boreham (1965), Hall (1972, 1974) California;
and Corredor (1975) Washington.
Northeastern U.S. works include: Johannsen (1903), Leonard (1926),
Metcalf (1932), DeFoliart (1951), Stone and Jamnback (1955), Jamnback
and Stone (1957), Jamnback (1969), Pinkovsky (1970), and, reporting the
unusual find of a South American black fly in the U.S. Wygodzinsky (1973)
- New York; O'Kane (1926) New Hampshire; Frost (1949) Pennsylvania;
Dimond and Hart (1953) Rhode Is.; Sutherland and Darsie (1960a and b)
- Delaware; Stone (1964) Connecticut; Holbrook (1967) Massachussetts;
Eckhart and Snetsinger (1969) Pennsylvania; and Crans and McCuiston
(1970a) New Jersey.
In the southeastern U.S. Tucker (1920) gave accounts of personal
experiences with black flies and reported two species, S. pecuarun and
S. meridionale, from Louisiana. Jones and Richey (1956) conducted a
survey of the black flies of Jasper County, South Carolina, and discussed
the biology, ecology and relationships to Leucocytozoon in turkeys of
one Cnephia (pecuarum) and seven Simulium species (congareenarum, deconk ,
jenningsi, slossonae, tuberosum, voenuatum and undescribed species).
Snow et al. (1958) reported on the ecology, habits and distribution of
black flies occurring in the Tennessee River Basin and recorded species
in the three genera: Chephia mutate2), Prosimutium~ (hirtipes, magnum,
plus undescribed species), and Silmu~ium (decorum, fibrinflatum~, jenningsi
ap. group, merid~ionate, pictipes, tuberose, veuan~tum, verecundum,
vittatum and three undescribed species). Snoddy and Hays (1966) mentioned
that eleven species of Simuliidae were captured at one location in
Alabama using a New Jersey light trap modified for daylight use by
removing the light and substituting,as an attractant, carbon dioxide
gas dispensed at .45 kg (1 lb)/hr. Stone and Snoddy (1969) present
distribution records, descriptions, and some biological information for
28 species of black flies discovered or expected to occur in Alabama.
Garris et al. (1975) present the seasonal distribution of 7 species of
Simulium collected from streams in Sumter Co., South Carolina, and
mention observations of black flies feeding on turkeys and captured in
a CO2-baited trap. Snoddy and Beshear (1968), and Snoddy (1971, 1976)
describe and give some facts on the biology of three new species
(S. tcaodiumi, podocstem~i and Zakei) in the expanding species list of the
former S. jenningsi group.
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
Eggs. Ussova (1961) describes most black fly eggs as between .24
and .33 mm long and irregularly triangular in shape with rounded corners;
the eggs of some Cnephia species, however, are said to be larger and
elongate elliptical. Golini and Davies (1975) found black fly eggs were
.228 mm long and .139 mm wide. Davies and Peterson (1956) examined the
eggs of four genera (Gymnopais, ProsimuZium, Cnephia, Simutium) and
found no external sculpturing. The eggs are light in color when just
laid and darken as the embryos develop. Davies and Peterson (1956)
found that the eggs of Gymnopais and Prosimulium were the largest and
those of ProsimuZium and C~ep~hia were the narrowest of the genera checked.
Black fly eggs are often found attached to vegetation and rocks in a
swift flowing stream or river or lying free on the stream bottom.
Tarshis (1968) conducted experiments on and reviewed accounts in the
literature of desiccation and the overwintering of black fly eggs and
concluded, as Wu (1930) had done earlier, that only those eggs which
are kept moist, by damp stones or underground springs, etc., in ap-
parently dry stream bottoms and other situations are able to remain
viable. Tarshis (1968) found that freezing eggs of a number of species
at 0 to-700C killed the embryos, while he was able to maintain moistened
eggs alive for 424 days at 2-90C. Kurtak (1974) found that eggs of
SimuZ~ium pictipes, recovered while encased in ice from rock crevices
along a stream, hatched in the lab after three days in 100C flowing
water. In northern areas eggs of many black fly species hatch when the
water temperature reaches around 80C (Carlsson, 1967). Raybould and
Grunewald (1975) found eggs of the K~ibwezi form of S. down2osrum watch at
200C four days after oviposition. Tarshis (1968) indicates normal egg
hatch for Maryland black fly eggs occurs within one to five days.
Larvae general morph~ology and development Black fly larvae are
usually between 4.5 andl 10 mm long when mature, club-shaped with a wider
posterior end, and possess a pair of multiple-rayed cephalic fans on
the anterior end, an unsegmented ventral proleg in the thoracic region,
and a circlet of anal hooks posteriorly. Dark cephalic head spots mark
the origins of cephalic muscles. Tarshis (1968) found that larvae
hatched from eggs in the lab after 7-38 days at 1000 and 1-5 days at
20-250C. Larvae developed to pupae in 18-50 days at 100C and 11-29 days
at 20-250C. Cameron (1922), Dalmat (1955) and Reisen (1975) found six
larval instars in the black flies they studied. Johnson and Pengelly
(1970) observed S. rugglesi to pass through seven larval instars and
Fredeen (1975) mentioned a seventh and final instar for S. aration.
Craig (1975) distinguished nine larval instars in Tahitian species of
Larval habitat. Larvae frequently attach by their posterior hooks
onto a patch of silk secreted by their large, looped salivary glands.
The silk is often applied to the same substrate on which the eggs are
found. Larvae may drift a short distance tethered to the old substrate
by a silk thread (Tarshis and Neil, 1970). A larva may also move in a
looping, geometrid fashion from silk patch to silk patch alternately
attaching its proleg then its anal disc (Dalmat, 1955). Elliot (1971)
reported that black fly larvae can actively migrate upstream as well as
passively downstream. Clean, smooth items free of algae or slime
are preferred attachment substrates for the larvae and larvae are
often located on such objects in stream sections where the current
is increased by a partial obstruction (Dalmat, 1955; Carlsson, 1967).
Larvae show positive phototaxis and colonize light substrates faster
and more densely than dark substrates (Carlsson, 1967). Unusual
attachment sites include the phoretic associations of black fly larvae
on prawns, mayfly nymphs, crabs, and other arthopods reported by
Disney (1971, 1973, 1975) and thle attachment of young larvae of S.
damnosum to older larvae of the same species (Burton, 1971). Carlsson
(1967) in Sweden found the greatest concentrations of larvae in flows
80 to 120 cm /sec although some species preferred 40
cm/sec currents. Cariase (1962) in the Philippines found immature black
flies concentrated where the water velocity was .5-.86 m/sec (1.63-2.83
ft/sec). Wu (1930) reported that simulfid larvae remained well estab-
lished at a velocity of 1.83 m/sec (6 ft/sec). Rohdendorf (1974) sug-
gested that the closed respiratory system of black fly larvae resulted
after invasion of the swift flowing well oxygenated habitat typical of
most simuliids. Tarshis (1968) found that black fly larvae could not
live more than eight hours in still water. Anderson and Shemanchuk (1975),
however, report they transported black fly larvae for several days in
shallow, non-agitated, ice-chilled water with little mortality. Wu
(1930) found approximately equal amounts of dissolved oxygen in quiet
and turbulent sections of black fly streams and after a number of ex-
periments concluded that black fly larvae have a definite requirement
for current to lessen sedimentation and supply adequate food, not because
of improved oxygen conditions. Crans and McCuiston (1970b)found larvae
Ln permanent rivers and streams, temporary creeks and flowing roadside
ditches. Dalmat (1955) and Lewis and Bennett (1975) reported finding a
few live larvae in situations where the flow was so slow that silt and
mud present prevented the larvae from anchoring to any fixed object.
Van Someren (1944) reported S. ruficorne larvae from small pools and
footprints in sandy river beds in Somaliland. Certain species are
typically found in limited type's of lotic habitats: S. arcticum in
large rivers (Cameron, 1922), S. pictipes at the outflow of damns and on
flat bedrock (Snow et al., 1958) and S. cohraceum in very small streams
about one meter (a few feet) wide and a few centimeters (a few inches)
deep (Dalmat, 1955). Travis and Vargas (1970) found that in contrast
to the clear mountain streams, the lower, slower Costa Rican streams
most polluted with sewage and garbage had the greatest larval and pupal
populations. Cariaso (1962) reported that black fly larvaeweredis-
couraged from breeding in water polluted with nitrogenous and human
wastes. Carlsson (1967) indicates moderate pollution increases organic
drift and is good for larvae but heavy pollution clogs the cephalic
fans. Streams in Wisconsin carrying large amounts of eroded soil par-
ticles and other detritus were observed to be poor habitats for most
black fly larvae due to feeding interference (Anderson and Dicke, 1960).
Larvae feeding structures, behavior, and food. Davies (1974)
and Craig (1974) describe the evolution and development of the important
larval feeding structures in many species, the cephalic fans, and men-
tion how these lateral palatal brushes which are well developed filter-
ing devices in most black flies are apomorphically absent in later
instars of Gyrmnopais and Zainnia and modified into raking structures
in Crosetia species. Chance (1970) stated that particles from about 1
to 350 microns in diameter, the majority in the 10 to 100 micron range,
were ingested by four filter feeding species she studied. Larvae were
found to attach within 10 centimeters of the water surface, extend their
bodies into the flow and alternately open and close the fans combing
particles from the fans when they were closed into the cibarium where
a bolus was formed. Mulla and Lacey (1976) report larvae, with their
heads pointed downstream, attach to silk deposits with their anal hooks,
rotate the body 900 to 1800 and open the cephalic fans to strain matter
from the flow. Chance (19703) suggests that the horizontally operating
mandibles, especially in non-filtering, grazing forms like Tannia,
scrape organic food off the substrate. Rubtsov (1974) discusses the
outer sclerite of the labium the submentum or hypostomum, an anteriorly
serrate structure bearing setae which lies just cephalad of the
gular notch and mentions that it is a tactile organ, is important in
producing the larval silk strands, and that it may serve to scrape food
from the substrate. In species which lack fans the submental teeth
sometimes acquire a spatulate shape.
Jobbins-Pomeroy (1916) reported Euglenaand Spirogyra were important
food items for black fly larvae but Cameron (1922) found diatoms formed
the main components of the food. Anderson and Dicke (1960) found
diatoms, other algae, and considerable inorganic material in the in-
testinal contents of black fly larvae. Carlsson (1967) listed as larval
food bacteria, plankton, plant and animal parts, pollen and aerial
fallout. Spring flooding leads to large outbreaks of black flies when
rising water temperatures trigger synchronous hatching of many black
fly eggs. The rising stream and river waters make more substrate avail-
able for larval attachment and increase organic debris and hence food
available for the larvae. Bacteria filtered out by S. underhilli led
to speculation of using simuliid larvae as indicators of pollution
(Snoddy and Chipley, 1971). Reisen (1974) found that larvae remove
bacteria and organic and inorganic particulate matter. Young larvae of
Simulium species in nature fed at a faster rate than did older larvae
as indicated by the time necessary to void a plug of dye particles
(Mulla and Lacey, 1976). These authors also found that at a lower
temperature (12.80C 550F) a longer time (35-55 min) was necessary to
eliminate a particulate plug from larval guts than at a higher tempera-
ture (300C = 860F, 20-30 min).
Pupae. Pupae attach to the same substrates as the larvae. The
larvae form silken cocoons in which pupation occurs. Hinton (1958)
mentions a pharate pupa stage where pupal characters are visible within
the last larval skin but the larval mouth parts are still articulated
and feeding continues up to when this "pupa" begins spinning the cocoon.
The cocoons may be thin and rudimentary (Rrinnia), shapeless, irregular
masses of silk (Prosim~utium and some Cnephia), sturdy, coarse cocoons
(Sinuium pictipes) or tapered, slipper-like finely shaped and tightly
woven cocoons (most SimuZium) (Stone and Jamnbackc, 1955). Members of
the Simuliumn subgenus Phosterodoros have cocoons with forward edges that
are convex in profile and have a large opening anteriorly on each side
of the cocoon (Stone and Snoddy, 1969). Field and Law (1961) describe
what Underhill (1944) illustrated for S. jenningsi (as S. nigroparvum),
that is, sexual dimorphism in the cephalic plate of simuliid pupae. The
cephalic plate, a sclerite that begins between the bases of the antennal
sheaths and extends over the cephalic area, is longer and narrower
in the male than it is in the female pupa. The pupae usually face down-
stream and are bordered on each side by tubular filaments which vary in
number and shape according to species and aid in respiration. Cameron
(1922) observed that the number of pupal filaments in S. arcticumt varied
from the usual 12 to 13 or 11 and that one set might vary while the
other group on the same pupa might bear the more typical number of
filaments. Coscaron and Wygodzinsky (1973) mention the variation in
the point of bifurcation or petiole lengths of the respiratory filaments
of different specimens within the same species.
Adults emergence. Tarshis (1968) reported that pupal develop-
ment of three species of Simulium took 1 to 5 days at 20-250C and that
at 100C it took 21 days following pupation before adults began emerging.
Davies et al. (1962) describe how, at emergence, after gas fills the
skin of the submerged pupa, the adult ruptures the pupal exuvium, pulls
itself through a T-shaped opening and quickly floats to the surface in
a bubble of gas (also see Hannay and Bond, 1971, section below). Carlsson
(1967) found that a heavy silt cover on pupae prevented adults from
emerging. Adults may easily be reared from pupae placed on moist filter
paper in petri dishes (Hannay and Bond, 1971; Sutcliffe and McIver,
1974). Wenk (1965) found males emerge sooner from pupae than do females.
Disney (1969) found three species of Simulium in Africa pupate by day
and may prolong pupation to avoid emerging at night. On warm days
eclosion occurred early in the morning, on cooler days eclosion peaks
occurred during the late morning and mid-day and under artificially cold
days emergence was shifted to the late afternoon.
Adults appearance and morphology. Adult black flies rarely ex-
ceed 5 mm in length (Dalmat, 1955). Male and female black flies are
hump-backed in appearance. The males are holoptic, usuallydarkerand
morevelvety than females and often bear a shiny pair of anterior lateral
spots on the scutum (Davies et al., 1962). Costalization (i.e., a
strengthening of the anterior wing veins and a decrease in wing venation
in the posterior portion of the wing), shortening of the legs, evolution
of the calcipala and pedisulcus, increase in head size and shortening of
the abdomen with reduction of tergites and sternites are considered
apomorphic developments (Rohdendorf, 1974; Rubtsov, 1974). Hungerford
(1914) in discussing the anatomy of Simulium vittatumt adults noted that
black fly females possess a single spermatheca. Lewis (1957) presented
morphological information about adults of S. damnnsum and suggestions
on dissection techniques. Bennett (1963b)found the shape and appearance
of the salivary glands of adults valuable in differentiating species.
Hannay and Bond (1971) found raised cylindrical buttons, each with a
wax filament, between the macratrichia on black fly wings and suggest
these structures may aid the adult in keeping its wings dry during
emergence. Sutcliffe and Melver (1974) describedcleaning hairs and
combs of cuticular teeth on the metathoracic legs and cleaning hairs
on the prothoracic legs which are used to clean the wings and head
Adults attraction and trapping. Wirth and Stone (1956) indicate
that male black flies are readily collected at light traps. Snow et al.
(1958) captured adults at light, on vegetation, in cars and biting
mammalian hosts. Fallis and Smith (1964) succeeded in using ether
extracts of birds as attractants for simuliids. Anderson and DeFoliart
(1961) used a variety of caged birds to attract ornithophilic black
flies to traps in Wisconsin. Golini and Davies (1971) found that fe-
males of S. venustum fly upwind to a carbon dioxide source and cease
flying upwind if the CO2 source is turned off. Snoddy and Hays (1966)
and DeFoliart and Morris (1967) made use of the attractancy of CO2! to
capture simuliids. The silhouette of a target rather than the reflec-
tance is more important in attracting black flies (Gillies, 1974).
Peschken and Thorsteinson (1965) state that black flies are more at-
tracted to stationary targets than to moving targets and simuliids show
less discrimination between form and shape of three dimensional objects
than some other hematophagous insects. Bradbury and Bennett (1974a)
found bloodseeking black flies in the genera Prosimulium, Onephia, and
Simuliwn were more attracted to black, blue, and red than white or
yellow, especially when the colors were of low visible reflectance.
Thorsteinson et al. (1965) describe a tripiod canopy-type Manitoba fly
trap which uses a large black sphere target to make the trap visually
attractive to bloodsucking diptera. Bradbury and Bennett (1974b) found
that black flies could distinguish between targets based on their color,
largely independent of carbon dioxide flow rates. Carlsson (1967) stated
lakes, shoals, bogs and swamps act as "collecting mirrors" for female
black flies seeking ovipositioning spots. Female black flies seeking
oviposition sites oviposited more on green and yellow colored plastic
strips in a stream than on red, purple, white or black strips (Golini
and Davies, 1975). The Malaise trap, a final valuable collecting device
for black flies and other insects,was conceived as a passive or random
flight interception trap;however there is some evidence that the degree
of color contrast between the trap and background vegetation is an
attractance factor (Roberts, 1970 and 1972).
Adult mating. Females of C. de~cotensis have mature eggs at the
end of the pupal stage and copulate with males on damp rocks soon after
emergence. In general, however, simuliids can mate any time from
emergence to oviposition time (Davies and Peterson, 1956). Davies and
Peterson (1956) were unable to induce male black flies to mate by broad-
casting the sound of the female wing beat frequency. Compared to males
which do not form mating swarms, males with mating flights have larger
eyes and enlarged upper facets. Snow et al. (1958) observed that males
of S. pictipes flew upside down with their abdomens curved upward, con-
tacted females as they flew by overhead, settled to a solid substrate,
and copulated venter to venter with the female situated uppermost. The
presence of about one male for every two hundred females of onephia mutata
is considered evidence for parthenogenic reproduction (Davies and Peterson,
1956). Downes (1965) reports Prosimulium .ursinum is a maleless species
in which the females do not even emerge from the pupa but disintegrate
shedding the eggs into the stream.
Adult feeding. Black flies have been collected from flowers
(Davies and Peterson, 1956). Hocking (1953) stated that black flies and
other northern biting flies obtain the energy for flight almost entirely
from floral nectar. Lewis and Domoney (1966) reviewed the importance of
sugar feeding on bloodsucking, autogeny, and parasite development and re-
ported finding glucose, sucrose, fructose and other sugars in the crops
of 101 wild caught Simutium. There are scattered reports of black flies
being attracted to and feeding on cold-blooded animals (Hagen, 1883;
Jobbins-Pomeroy, 1916; Smith, 1969), but of more importance are their
ornithophilic, mammalophilic, and anthropophilic bloodsucking habits.
Onephia dacotensis, Gyrmnopais hotoptious, Tw~innia ti~bbesi and a few
other black fly species acquire sufficient nutrients during the larval
stage to produce eggs and do not suck blood (Davies and Peterson, 1956;
Shewell, 1957). Rohdendorf (1974) suggested that limited larval food
stimulated the adult female black flies to hunt and feed on vertebrates.
Cameron (1922) mentions the tenacity and persistence black flies ex-
hibit when feeding and states that once the mouth parts are securely
inserted into the skin the insect is not easily disturbed. Ussova (1961)
states that the mandibles pierce a host's skin, the maxillae with re-
curved teeth anchor the proboscis in the skin, and alternating actions
of the cibarial and pharyngeal pumps suck the blood into the esophagus.
James and Harwood (1969) indicate simuliids are telmophages or pool
feeders which lacerate blood vessels with the toothed, transversely
operating mandibles and vertically operating maxillae. Davies and
Peterson (1956) record a wide, natural host range for some species such
as S. venustum duck, crow, heron, deer, and human. Others such as
P. hirtipes (now three species) and S. decorum which feed on? mammals in
the field fed on birds when placed in vials on the avian species in
the lab. Yang and Davies (1974) examined the salivary glands of three
black flies and found an anticoagulant factor which keeps the blood
fluid for movement into the gut. These authors also report an agglutin
factor in flies at least twelve hours old.
Adult egg development and egg laying. Cameron (1922) found
development of the ovaries of female black flies followed a successful
blood meal. Davies and Peterson (1956) describe females of several
species with weak teeth or only hairs on the mandibles and maxillae. These
flies do not suck blood and emerge with already ,mature eggs. Davies
and Peterson (1956) indicate eggs develop five to twenty-one days after
emergence with the longer time involving a prolonged blood meal search
under natural conditions. Immature and mature eggs are found together
in the ovaries which suggests at least two ovarian cycles (Cameron,
1922). Females may survive long enough for three batches of eggs but
few probably live this long in the field (Davies and Peterson, 1956).
Cameron (1922) reported that the eggs of Simutium simile (=S. arctioum)
are oviposited on rocks in large cake-like masses embedded in a soft
gelatinous matrix, Davies and Peterson (1956) mention Prosimulium and
Cnephia species which deposit eggs freely while flying down or across
a stream. Stone and Jamnback (1955) indicate S. vittatum lays strings
of eggs in a gelatinouls matrix. Golini and Davies (1975) in Canada
found that female black flies settled at the water line on trailing
cattail leaves and deposited large (16 cm x 2 em) irregular egg masses
one to five layers deep in a gelatinous substance. Davies and Peterson
(1956) found simuliids oviposited an average of 200 to 500 eggs per
female while Golini and Davies (1975) report an average of 417 eggs
for each female.
Adults life cycle, longevity, range, and general activity.
Tarshis (1968) reports a 21 to 25 day period for mixed cultures of five
Simuliaum species to develop into adults after eggs were placed in aquar-
ia. Raybould and Grunewald (1975) found developmental time from egg to
adult ranged from 18 to 50 days for African black flies.
Dalmat (1955) in Guatemala used colored dyes and the release-
recapture technique and found marked female black flies survived as
long as 85 days and traveled as far as 9.7 miles. Bennett (1963a) tagged
females of S. rugglesi by feeding them on ducks injected with a phos-
phorous-32 solution and recovered labelled flies up to 28 days later and
9.6 km? distant following watercourse paths. West-et al. (1968) exposed
black fly larvae and pupae to 32P-treated water and recovered radioactive
flies as far as 33.5 km (20.8 mi) distant. Bennett and Fallis (1971)
found S. eurycdniniculum flew up to five miles from the release point
and report the average life span of the females was at least two to
three weeks. Hocking (1953) summarized published records of flight
ranges which reached a maximum of 145 km for S. reptans columbaczen~se.
Wellington (1974) reported frenzied activity in black flies which
was apparently correlated with barometric pressure changes as a storm
system approached. Cameron (1922) observed swarms of male black flies
or males and females on warm, cloudy days with rain threatening or
falling gently. Carlsson (1967) reports that rapid changes in tempera-
ture, air pressure, and light seem to increase the activity of all
simuliid species but he also mentions even a light breeze reduced black
fly activity considerably. Hocking (1953) found flight was continuous
in Simulium species down as low as 12.80C (550F).
Concerning nocturnal behavior Dalmat (1955) observed that black
flies in Guatemala move down to the base of plants as the sun sets and
can be captured at night using a lantern and sheet when the insects
emerge from their resting sites near the ground. Wolfe and Peterson
(1960) describe climbing trees and sweep netting twenty-five feet off the
ground to capture black flies at night in Quebec. A4t dawn Wolfe and
Peterson observed simuliids flying down from the canopy. Recently in
India researchers used light traps and found that black flies were active
throughout the night with a peak of flight activity occurring around
midnight (Datta and Dasgupta, 1974).
Lab colonization. Black flies have been collected in nature in all
stages and have been brought to the lab for experiments. Tarshis (1965a)
describes techniques for collecting and shipping viable black fly eggs.
Tarshis (1965b) and Tarshis and Adkins (1971) discuss collecting
large numbers of black fly larvae on artificial cloth substrates and
mention techniques for transporting larvae in aerated containers.
Tarshis (1971) discusses rearing black fly pupae to adults in individual
brass strainer cloth cylinders connected to aquarium air stone units.
Tarshis (1972, 1973) describes field collection, laboratory rearing of
immatures to adults, and successful feedings of females of C. ornitho-
philia on ducklings in the lab.
Wenk and Raybould (1972) point out that colonies of insects aid
critical studies on their biology, provide adults free from infection
for transmission work and can provide abundant material for investiga-
tions at times when natural populations are limited. Dunbar (1969)
mentions that hybridization experiments supported by colonies of flies
and sufficient knowledge of mating requirements can provide valuable
information concerning the species status of insect forms. Dalmat (1955)
found carbon dioxide stimulated feeding and oviposition and succeeded in
inducing 40% of 65,000 S. ochraceum, S. callidum,; and S. metaieum to
feed and 20% to oviposit but no eggs developed into larvae. Pield et al.
(1967) found that females of S. vitbatum confined in a vial bit man,
quail, and rabbits. By impaling a male on a minute and brushing the
male genitalia against the genitalia of the female coupling could be
achieved. No female of S. vittatum which mated survived to oviposit
and eggs deposited by other, unmated, females in the artificial flows
available failed to hatch. In the laboratory Wenk (1965) successfully
reared, mated, blood fed and achieved oviposition of viable eggs with
Boophthora ery~throcephala. Wenk and Raybould (1972) working with the
Kibwezi form of S. damnrosum likewise reared, mated, blood fed and ob-
tained viable eggs from female black flies in the lab. Mating, which
was also achieved with a member of the S. bovis complex, occurred in
partially lighted emergence cages and was confirmed by the presence of
sperm or spermatophores. Raybould and Grunewald (1975) review the litera-
ture on lab colonization of black flies and mention difficulties confront-
ing researchers: inducing mating, inducing females to feed on blood suf-
ficiently and consistently, erratic viability or hatchibility of eggs
oviposited in the lab, and removing wastes from immature rearing setups.
Although a few black fly species have been induced to complete every
stage of their life cycle in the lab, they are still not considered to
be successfully colonized.
Jamnback (1973) describes the black fly bite reaction: a hemor-
rhagic spot which develops into an itching wheal; sensitive individuals
may suffer headache, fever, nausea, glandular enlargement, and bites
around the eyes may cause swelling that results in obscured vision.
Loomis et al. (1975) report black flies cause considerable irritation
and tissue damage to horses' ears, head, neck, and belly,producing
wounds and papules. S. archiewn, C. pecwuarn, and S. columbaczense are
described as major pests of livestock (Jamnback, 1973). Fredeen (1974)
mentions black fly outbreaks during 1944-1947 in Saskatchewan which re-
sulted in the deaths of 1100 farm animals including cows, horses, hogs,
and shorn sheep. Fredeen (1975) reports a S. arcticum outbreak in 1972
along the N'orthern Saskatchewan River that killed at least 18 cattle.
Black flies transmit Leucocytozoon parasites to ducks (Fallis and
Bennett, 1966), trypanosomes to ducks (Desser et al., 1975), and
Leucocytozoon parasites to turkeys (Skidmore, 1931; Noblet et al., 1972)
(see also the Leuc-ocytozoon smithi sections in the Literature Review and
Results and Discussion below). Sudia et al. (1975) mention that epi-
demic VEE virus has been isolated from Simuli~um species in Colombia and
although biologic transmission has not been proven, mechanical trans-
mission by contaminated mouth pprts may be possible for at least 72
hours. Travis et al. (1974) report a finding of vesicular stomatitis
virus in black flies in Colombia. Eastern encephalitis virus has been
isolated from a pool of 100 unengorged S. meridionale which indicates
this species may biologically transmit the disease (Anderson et al.,
Steelman (1976) reports monetary losses in cattle herds due to
worm nodules caused by simuliid transmitted Onchocerca gutterosa amount
to $500,000 each year in Australia. Onchooerca voloulus,the causative
agent of blinding filariasis in mantis vectored by black flies and has
been reported from Africa, Yemen, Mexico, Guatemala, Venezuela, Colombia,
and Surinamf; indigenous cases may also occur in Brazil (World Health
Organization, 1971; Travis et al., 1974; Raybould and Grunewald, 1975).
i~crofilaria of 0. voloulus previously were observed in the eyes and
skin, however, Anderson et al. (1975a) found microfilaria in the urine,
blood, and sputum of persons treated with diethylcarbamazine. Diurnal
periodicity of 0. volvulus microfilaria in the skin has been shown to
correspond with the peak feeding periods of important black fly vectors
in Guatemala and Africa (Anderson et al., 1975b). Duke et al. (1975)
found the transmission potentials of 0. volvulus along breeding rivers
for S. d'awnosum in Africa so high that no communities could survive there.
Physical control. Impoundage of rivers and the creation of reser-
voirs together with planned periodic cutoffs of discharge water has
discouraged or eliminated black fly breeding along many stretches of
rivers (Snow et al., 1958). Removing debris such as planks, tree limbs,
trailing vegetation and other potential substrates for black fly larvae
can help control s~imuliid population levels in streams (Jamnback, 1973).
Chemical control. Carestia et al. (1974) reported that serial ap-
plications of malathion, DibromR or Dibrom plus a repellent for
adult black fly control were unsatisfactory. High winds, dropping
temperatures and decreasing daylight interfered with adult fly activity
and chemical effectiveness. Rapid reinfestation of adults from areas
just outside the treated zone is another reason why most control efforts
have been aimed at immature rather than adult black flies.
Fairchild and Barreda (1945) observed the effectiveness of DDT as
a black fly larvicide. Davis et al. (1957) discussed early attempts at
evaluating the larvicidal properties of parathion dripped into streams,
dieldrin applied by air, and DDT delivered by various ground techniques
as well as by airplane. Evidence of the persistence of DDT in non-target
organisms and decreased susceptibility of black fly larvae from streams
with a history of DDT treatments stimulated a switch to less persistent
materials (Fredeen et al., 1971; Jamnback and West, 1970). Travis et al.
(1970) in field trough tests found Dibram was outstanding in producing
mortality of S. pictipes larvae at .5 ppm and 1 ppm concentrations and
stated Dibrom, GardonaR and CiodrinR deserved more practical stream
tests.' Jamaback and Frempong-Boadu (1966) found chemical formulations
are most effective which give uniform distribution of the insecticide
in the water and have a specific gravity of slightly less than 1.0 to
keep the toxicant near the surface where most of the black fly larvae
are located. AbateR (20%) in Panasol plus .5% Triton X-161 (.998
specific gravity) applied by aircraft eliminated larvae up to .8 km
(.5 mi) downstream from the treatment point. Pelsue et al. (1970) chose
Abate as a black fly larvicide due to its low mammalian, avian, and fish
toxicity and achieved 100% control of larvae in three locations
with .5 ppm delivered for 60 minutes. Detachment of larvae occurred
within 24 hours and reinfestation was noted in 15 to 60 days. Kissam
et al. (1975) reported 2% Abate Celatom granules delivered at 91 g AI/
.4 ha monthly reduced larval populations t~o zero and the populations
only slowly built back up during the two weeks following each applica-
Fredeen (1974) mentions that methoxychlor, another black fly larvi-
cide, is minimally toxic to vertebrates and methoxychllor and its
metabolites are not concentrated in fish or other aquatic species. In
Canada in 1969 a 15 minute injection of .2 ppm methoxychlor emulsifiable
concentrate caused the disappearance of 96% of the S. arcticum larvae
32 km downstream. Fredeen (1975) reported that a 7.5 minute injection
of methoxychlor at .6 ppm killed 100% of the older black fly instars 40
and 80 km below the treatment area. Younger black fly instars which are
more susceptible were depleted by 96% at a site 161 km downstream from
the injection. Fredeen et al. (1975) indicate that methoxychlor adsorbs
to suspended solids in the water and may act selectively against the
filter-feeding black fly larvae. Following a methoxychlor application
Wallace et al. (1973) noted increased drift but no eradication of non-
target organisms and Fredeen (1975) found that non-target organisms
repopulated more densely than before the treatment.
Investigations of the effect of insect developmental inhibitors on
black flies indicate that significant reduction (75-100%) of adult
emergence can be attained in the lab with at least three black fly
species with AlcosidR at concentrations between .001 and 1 ppm (McKague
and Wood, 1974; Dove and McKague, 1975). Frormmer et al. (1975) reported
that in field evaluation tests in New York DEET-treated light mesh
jackets were effective in reducing landings per five minutes from 404
on controls to 1.2 on personnel with treated jackets.
Biological control. Cameron (1922) found that a fish of no food
value called the common sucker fed on S. simile (=S. arcticum) larvae
and pupae. Peterson and Davies (1960) review the predators of black
flies which include adult Empididae and Dolichopodidae, Tendipedidae
larvae, Trichoptera, Formicidae, Odonata -adults and naiads, and spiders.
Sommerman (1962) reported empidid larvae fed on black fly larvae.
Snoddy (1968) found the solitary wasp Oxrybelus emarginatum to be a
predator of adult black flies. Peterson and Davies (1960) and Chutter
(1972) report cannibalism among simuliid larvae. Carlsson (1967) in-
dicates predators are unlikely to serve well as man-manipulated biologi-
cal control agents though the tricopteran genera Hiydropsyche and
R~hyacophila might give good results.
One of the earliest accounts of parasitism of black flies is that
.of Strickland (1911) who reported a mermithid nematode that retarded
the development of the larval histoblasts and a sporozoan (microspori-
dian) which caused distorted and swollen larvae. Jamnbackc (1973) states
that eighteen microsporidia have been described from black flies.
Spores are ingested by the larvae, sporoplasm invades suitable host
cells which are often in the fat bodies, multiplication and the pro-
duction of many spores occurs, and the host larvae commonly die. Black
fly pupae and adults have been found infected and transovarian trans-
mission is common. Jamnback (1973) also mentions a fungus Coeiomycidiwn
simulii which occurs in black fly larvae and is usually fatal.
Davies (1958) reports microsporidians, mites, and mermithid nema-
todes as parasites of Canadian black flies and mentions that mermithids
are found in larvae, pupae and both sexes of the adults. From 15 to
60% of the females in emergence collections were found to be infected
with mermithids and females in oviposition swarms were also found with
mermithids. The infected females which attempt to oviposit may serve to
disperse the mermithids and introduce them to additional stream environ-
ments. Welch (1964) indicated that the mermithid life cycle began with
the consumption by black fly larvae of the mermithid, usually in the
infective first juvenile stage. Molloy and Jamnback (1975) observed
direct penetration of the black fly larval cuticle by preparasitic
juveniles of Neomesomermis fluiminais, a mermithid which infects at
least fourteen species of black flies in three genera. Exit of the
parasite from the host is almost immediately fatal to the black fly.
Anderson and DeFoliart (1962) report 49-93% parasitism by Isomermis and
Gastromermis in one black fly species in Wisconsin. As a step toward
the mass rearing of these biological control agents Bailey et al. (1974)
discuss techniques for the mass collection of mermithid postparasites
from field collected black fly larvae.
Batson et al. (1976) reportedatiridescent virus from black fly
larvae in Wales but could associate no major fluctuations in the black
fly population with the presence of the infected larvae.
Florida Ecological Habitats
Florida is a peninsula that lies between 80 and 88 degrees west
longitude and in the same latitude belt as the Sahara, Arabian, and other
large deserts. Exclusive of the Keys, the State extends roughly 644 km
(400 mi) north and south along the peninsula and about the same distance
east to west along the north coast of the Gulf of Mexico. Florida covers
151,710 sq km (58,560 sq mi)'. The highest point in the State is 105 m
(345 ft) above sea level, at Lakewood in Walton County. From Orlando
south to Sebring another high section occurs and Iron Mountain near
Lake Wales is located about 91.5 m (300 ft) above sea level (Morris,
1973). The earliest geological horizon is the Ocala limestone formed
during the Eocene, 58 million years ago. Modern Florida made its first
appearance during the Oligocene, 36 million years ago, as a small island
about where the counties of Suwanee, Columbia, and Alachua are now
located and then rose as a peninsula during the Miocene (Byers, 1930).
There are three schemes which have been used to break Florida up
into sections according to: 1) vegetation; 2) geology; and 3) edaphic
factors. These schemes are presented to help illustrate the ecological
diversity in Florida, which an the one hand provides a range of habitats
for simulfid species and on the other hand restricts black fly breeding
to certain more suitable regions of the state. Byers (1930) divided
Florida into seven biotic areas based on dominant types of vegetation:
a tropical hammock strip from St. Lucie County through Dade County along
the east coast extending a few miles inland with cabbage palms, mahogany,
ironwood, papaya, epiphytic bromeliads, plus other plants; a grass swamp
area south and east of Lake Okeechabee; magnolia and temperate hammocks,
primarily along the east coast from Flagler County to Indian River
County and on the Gulf Coast from Wakulla through Hernando Counties -
these occur on well drained but moisture holding soils with magnolia,
holly, and red bay as dominant vegetation on the east coast and beech,
elm, sweetgum, hickory, and oaks dominant in the west coast hammocks;
a section of southeastern deciduous forest extending down into Leon,
Liberty and Jefferson Counties; tree swamps in Collier, Columbia, Baker
and Nassau Counties which include Big Cypress Swamp and other cypress
and also black gum swamps; pine flatwoods along the Gulf west of
Wakulla County, on the west coast of the peninsula from Pasco to
Collier Counties, and in the Clay County and Putnam County region -
these flatwoods occur on level, poorly drained land underlain by hard
pan which results in an acid soil upon which grow palmetto, grass and
pines; and, lastly, the southeast coniferous forest with long leaf,
yellow, and slash pines and saw palmetto on well drained uplands.
Cooke(1939) divided Florida into five physiographic areas: central
highlands, Tallahassee hills, western highlands, Marianna lowlands and
coastal lowlands. The central highlands, in the center of the peninsula
from Baker County and Columbia County south to Lake Okeechobee, contain
in their southern section thousands of lakes which in the north section
are accompanied by numerous streams. The Tallahassee hills and western
highlands are composed mainly of red clay with a relatively good number
of streams. The coastal lowlands usually lie less than 30 m (100 ft)
above sea level and include swampy areas such as the Florida Everglades
and marshy areas along the east and west coasts of the state. The
Marianna lowlands in Wlalton, Holmes, Washington and Jackson Counties
contain few permanent flows and consist mainly of swamps, flatwoods,
and sandy hills bearing pines.
Davis (1967) and Smith et al. (1967) present six major land resource
areas for Florida: a southern coastal plain; gulf coast flatwoods;
central Florida ridge; Atlantic coast flatwoods; southern Florida flat-
woods; and Everglades and associated areas. The southern coastal plain,
occupying the northern half of the state from Escambia County to Madison
County,is covered by forests of mixed hardwoods and pines on the lower
areas and forests of long leaf pines and oaks on upland clay or well
drained upland sand. The gulf coast flatwoods--a strip along the gulf
side of the panhandle from Escambia County through Citrus County, the
Atlantic coast flatwoods primarily in Columbia, Baker, Nassau andBrad-
ford Counties in the north, and the south Florida flatwoods occupying
most of the southern section of the peninsula outside of the Everglades
consist of pine flatwoods and swamp forests (pines, palmetto, herbs,
bay tree, laurel, gum, cypress) on poorly to very poorly drained and
marshy soils. The central Florida ridge is located on well drained
soil in the middle of the peninsula and includes forests of long leaf
pine and xerophytic turkey oak (now mainly planted in citrus) and hard-
wood forests of mixed evergreen and deciduous hardwoods on rich upland
soils. The Everglades are primarily sawgrass, M~ariscus jamaioensis, on
peat and muck soils.
Average January temperatures range from 10-130C (50-550F) in the
panhandle to 18-210C (65-700F) in the Everglades area. Average July
temperatures fall between 27-290C (80-840F) throughout the state. An-
nual rainfall averages 134.62 cm (53 in) (Raisz, 1964). June through
September or October is considered the rainy season in Florida (Byers,
1930) however Berner (1950) indicates that in the northwestern area
rainfall is more evenly distributed throughout the year. There are
seventeen first magnitude springs in Florida (Morris, 1973). Despite
the heavy precipitation there are relatively few surface flows since
much of the water moves in underground channels in the underlying lime-
stone (Raisz, 1964). The vast majority of true streams and rivers are
found north of L~ake Okeechobee. Morris (1973) indicates there are 1,711
streams, rivers, and creeks in Florida with a total length of 16,989 km
Berner (1950) states that in Florida there are relatively few
intermittent streams. He divides permanently flowing streams in the
State into: sand-bottom creeks with loose shifting sand and little
vegetation; sand-bottom creeks with fairly loose sand and considerable
vegetation; silt-bottomed creeks with little vegetation; and silt-
bottomed creeks with considerable vegetation. Stagnant rivers occur in
south Florida where they serve primarily as drainage canals for the
Everglades and though there is considerable vegetation along the shores
of the rivers their fifteen to twenty-foot depths restrict plant growth
toward the middle. Larger calcareous streams contain water which rises
from springs, are basic in pH reaction, and in shallower sections are
lined on the bottom with Vazllisneria (eel grass), Sagittaria (arrowhead),
algae and other plants. Erving (1971) notes that the temperature for
Florida springs is about 220C (720F) year round. While noting springs
are alkaline Berner (1950) also indicates that marsh and swamp water can
be very acidic reaching a pH of 3.6 or below. He further adds slow
flowing rivers to his classification of Florida's streams and rivers
and included here? are the Suwanee, Apalachicola and other rivers which
have extensive drainage areas. Wood and Fernald (1974) list twenty-
eight major drainage basins and streams in Florida. The two largest,
the Apalachicola River Basin and the Suwanee River Basin drain 50,777
sq km (19,600 sq mi) and 25,984 sq km (10,030 sq mi), respectively,
some of which lies outside of Florida. Wood and Fernald (1974) indicate
that peak annual flows occur in eastern rivers in September and October
with buildups since June while more western rivers (Chipola, Apalachi-
cola) reach maximum flow during March and April. In northwesternl
Florida a continental weather pattern predominates whereas a more
tropical weather routine predominates in central and south Florida.
Beck (1965) recognized five stream types in Florida: sand-bottomed,
calcareous, swamp and bog, larger rivers, and canals. He points out
that due to velocity, substrate, and spring discharges many of these
types of flows can be recognized at different points along a single
watercourse and cites the Suwanee River which is successively a swamp
and bog stream, a sand-bottomed stream, a calcareous stream and a larger
river of mixed origin. Beck (1965) classifies swift flow in Florida's
streams as that velocity suitable for Plecopterawhile simuliid popula-
tions indicate conditions of moderate velocity. Below these two levels
the speed is considered low. A general progression is recognized with
the swifter portions of upper rivers marked by eroding limestone and
thick growths of moss, midpoints along rivers being sand-bottomed and
lower, slower portions of rivers containing deposits of finer materials
on the bottom. Beck (1965) lists the sand-bottomed stream as the most
common type of stream in Florida. The fauna consists of Hydropsychid
and Philopotamid caddistlies, simuliid larvae, Ple-coptera, Stenonema
mayflies, orthocladine chironomids, and Corydatis. In these sand-
bottomed streams the pH reaction is usually between 5.7 and 7.4 and
they display moderate to high color and moderate to swift velocity. The
bottoms consist of sand, leaf deposits, and limestone outeroppings, and
plant growth may be dense. Beck mentions that swamp and bog streams are
highly acidic (pH reaction 3.8-6.5), sluggish flows found in the coastal
lowlands and central highlands with origins in swamps and marshes.
Calcareous streams are cool, clear flows of spring origin with dense
growths of vegetation. These flows are alkaline (7.0-8.2 in pH), bear
large mollusc populations, are generally of low color, of variable
velocity and have sand, clay or limestone bottoms. The flows in the
larger river category usually carry considerable silt and are turbid,
have high and steep banks and bottoms of coarse sand and limestone.
They demonstrate a pH reaction of 6.5-7.4 and have few shallow places
and few aquatic plants. Beck (1965) mentions from about the area of
the St. Lucie Canal to south of Homestead no natural streams of any
consequence remain along the east coast of Florida and, instead, canals
are prevalent although in the past the Miami River had a swift flow and
rapids (Fairchild, 1976 personal communication).
The above discussion provides an introduction to the climatic,
edaphic, biotic and hydrologic conditions which are found in the State
During the 1880's Danilewsky observed parasites which were later
described and designated as species of Leucocytozoon and Haemoproteus
(Bennett et al., 1965). Hsu et al. (1973) list 67 species of
Leucocytozoon, one of which is reported from Meteagris gallopavo, the
domestic and wild turkey. Theobald Smith in 1895 first reported a
protozoan-parasite in the blood of turkeys in Massachusetts and Rhode
Island (Smith, 1895). Similar gametocytes were observed in the blood
of turkeys in France and named Haemamoeba smith (Laveran and Lucet,
1905). The following taxonomic scheme which shows the position of the
turkey parasite now known as Leucocytozoon smith was modified from
Aikawa and Sterling (1974) by crediting Sambon (1908) not Danilewsky
(according to Bennett et al., 1975) with the genus Leucocytozoon:
Phylum Protozoa Goldfuss, 1818, emend. von Siebold, 1845
Subphylum Sporozoa Leuckart, 1879
Class Telosporea, Schaudinn, 1900
Subclass Coccidia Leuckart, 1879
Order Eucoccida Leger and Duboscq, 1910
Suborder Haemosporina Danilewsky, 188i5
Family Leucocytozooidae Fallis and Bennett, 1961
Genus L~eucocytozoon Sambon, 1908
Species smith (Laveran and Lucet, 1905).
The following authors observed L. smith in turkeys in their respec-
tive states or countries: Atchley (1951), Bierer (1954), Jones and
Ricbey (1956) and W~ehr (1962) in South Carolina; Savage and Isa (1945)
and Bennett et al. (1965) in Canada; Volkmar (1929) in Minnesota and
North Dakota; Hinshaw and McNeil (1943) in California; Johnson (1945) in
Michigan; Kozicky (1948) in Pennsylvania; Skidmore (1931) in Nebraska;
Stoddard et al. (1952) in Georgia; and Byrd (1959) in Virginia. Volkmar
(1929) also lists L. amithi from Germany and the Crimean Peninsula and
Cook (1971) lists Asia. Travis et al. (1939) reported Leucocytoaoon
smith from wild and domestic turkeys in Florida as well as Georgia,
Missouri, Alabama and South Carolina. Simpson et al. (1956) and Forrester
et al. (1974) also reported Leu~cocytozoon smithi from Florida turkeys.
Solis (1973) exposed turkeys, ringnecked pheasants, chickens, bob'white
quail, pekin ducks and chukar partridges in an area where L~. snrithi
occurred and found that only turkeys were susceptible to L. smithi.
Fallis et al. (1974) and Aikawa and Sterling (1974) present in-
formation an the life cycle and structure of the intracellular parasite,
L. smith. Leucacotozoon belongs to the same suborder, Haemosporina,
as Plasmodiun and Hiaemoproteus and undergoes a similar life cycle.
Unllike PLasmodium, however, there is no eryrthrocytic schizogony and in
addition to the red blood cells being invaded by merozoites as in
Haemoproteus (and Plrsmrodiuw)lin L~aucocytozoon white blood cells are
also used as sites for gametogony (Huff, 1942; Fallis et al., 1974).
Gametocytes, in the case of L. smith, grow and split the host nucleus
in two (Sambon, 1908; Volkmar, 1929). Plasmodium species are tran~s-
mitted by mosquitoes and Haemnoproteus (Parahaemoproteus) species by the
Hippoboscidae and Ceratopogonidae (Fallis and Bennett, 1961a and b;
Bennett et al., 1974). Leucocytozoon, is known at present to be trans-
mitted only by black flies. A parasite of chickens which is vectored by
Culicoides was designated Akiba caulleryi by Bennett et al. (1965) but
Akiba has recently been considered as a subgenus of Leucocytozoon (Hsu
et al., 1973; Fallis et al., 1974). Gametocytes are ingested with the
blood of an infected turkey by a feeding black fly. Roller and Desser
(1973) observed diurnal periodicity with L. simondi where peak gameto-
cytemia corresponded with peak activity and biting periods of the vector.
It has been suggested that there is periodicity of the gametocytes of
L. smith in the peripheral blood and correlation with the periodicity
and feeding peaks of the vectors (Moore et al., 1974). Early in the
infection each gametocyte appears as a round body in the parasitized
cells; later the gametocytes deform each blood cell into the charac-
teristic spindle shape (Huff, 1942). Desser et al. (1970) found with
L. sim~ondi elongated cells resulted when merozoites developed in leuco-
cytes and round cells resulted from invasion of red blood cells. Macro-
gametacytes stain more darkly with Glemsa stain than microgametocytes.
Within minutes of ingestion the microgametocytes and macrogametocytes
escape from their host blood cells. The microgametocytes exflagellate
and syngamy occurs with the macrogametes. The zygotes formed grow into
elongate ookinetes within twelve hours after the initial blood ingestion.
Sections of flies have revealed ookinetes in the process of penetrating
the midgut of the fly (Fallis et al., 1974). Fallis and Bennett (1961a)
working with L. simonzdi detected only sluggish movement of ookinetes
and expressed doubt that this motion could facilitate penetration of
the gut wall of a black fly. Howard (1962) working with Plasmodium
gallinaceum1 observed no mobility or active penetration by zygotes.
Howard suggests a largely passive incorporation into the gut wall to
within 5 microns of the external basement membrane as the blood meal is
digested and the distended gut returns from a squamous cell configura-
tion to its original columnar cell morphology. Desser (1970) described
a pore, strengthening struts, microtubules and elongate convoluted
micronemes in the apical cap of the anterior end of L. simondi cokinetes
and suggested these structures might aid penetration through the peri-
trophic membrane and into the fly midgut epithelium. In as little as
48 hours the ookinetes develop into oocysts containing 50-100 sporozoites
just beneath the outer membrane of the gut. Sporozoites escape from the
oocysts and penetrate and collect in the salivary glands. Desser (1970)
describes structures on the anterior end of sporozoites of L. simondi
which may contain proteolytic enzymes and aid in entering the salivary
glands and, later, host liver cells. The sporozoites pass out the
proboscis of the black fly with the salivary fluids when the fly bites
another host. The sporozoites of L. sm~ithi invade the liver parenchymal
cells of the turkey and grow into hepatic schizonts. Huff (1942) re-
ports with other species of Leucocytozoon megaloschizonts in the heart,
spleen and other host tissues. Such schizonts are not normally observed
with L. smithi; however Siccardi et al. (1974) reported finding magalo-
schizonts in the kidneys of infected turkeys fed coccidiostat medication.
The schizonts by asexual multiplication (schizogony) produce merozoites
which enter red and white blood cells. Peters (1971) suggests that in
the related genus Plasmodium merozoite penetration may involve proteolytic
enzymes and be active or may be more passive with the merozoites being
engulfed or invaginated into the blood cells. The merozoites grow into
gametocytes in the blood cells and the cycle is completed. The prepatent
period or time elapsed between entrance of the sporozoites and appear-
ance of gametocytes in the peripheral blood is 10 to 16 days.
Symptoms associated with leucocytozoonosis include: loss of
appetite, emaciation, wheezing breathing, congested heart and lungs,
drooling, drooping wings, sitting on the hocks, enlarged liver and
spleen, anemia, impaired imrmunalogical responsiveness, and death (Wehr,
1962; Salsbury, 1971; Siccardi et al., 1974). Turkeys under 12 weeks
of age are often severely affected but mortality in older birds has
also been reported (Simpson et al., 1956). Many birds with high para-
sitemias appear outwardly normal. Birds that do not die may continue
to carry the parasite for as long as 14 years (Skidmore, 1931). Borg
(1953), Simpson et al. (1956) and others suggest that the pathogenicity
of L. smith has not been proven conclusively and that L. smith may be
an additive debilitating factor which when combined with other factors
such as blackhead, fowl cholera or leukosis results in bird mortality.
Stoddard at al. (1952), Skidmore (1931), Savage and Isa (1945) and
others however have observed flock mortality up to 75% where bacterial
cultures and other tests revealed L. smith as the sole disease organ-
ism present. Jones and Richey (1956) indicated annual death losses due
to Leucocytozoon in one county in South Carolina averaged 5%. In
1973,132 million turkeys were raised in the U.S. and farmers grossed
$934 million (Poultry Digest, April 1974; Agricultural Situation,
November 1974). In the absence of severe threats by disease this indus-
try will continue to grow as the public's demand for poultry and its
products like turkey rolls and t.v. dinners as an alternative to beef
A number of black flies have been incriminated as possible vectors
of L. smith. Skidmore (1931) incriminated S. occidentale (=meridionale)
as a vector of L. smith in Nebraska by grinding up a number of f-lies
that had fed on infected turkeys, injecting them in saline into clean
birds, and 12 days later observing gametocytes of L. smith in the tur-
keys' blood streams. Johnson et al. (1938) in Virginia exposed uninfected
turkeys to the bites of S. nigroparvum (=jenningsi) and obtained
L. amithi transmission. Underhill (1944) further substantiated this
work but Byrd (1959) was unable to infect clean turkeys by macerating
and injecting S. jenningsi that had fed on diseased birds. Wehr (1953)
by injection and Jones and Richey (1956) by fly-bite incriminated
S. skossonae as a vector of L. smith. Byrd (1959) succeeded in
infecting turkeys with L. smith by grinding and injecting females of
Prosimulium hirtipes that had fed on infected birds. I have seen no
further follow up of this work and Fallis et al.(1974) do not list any
member of the former P. hirti~pes complex as a vector of L. s ithi.
Noblet et al. (1972) ground up in saline and injected infected females
of S. congareoenruAm into turkeys and incriminated this species as a
suitable or possible vector for L. smith. Savage and Isa (1945) and
Anthony and Richey (1958) have reported leucocytozoonosis in the sup-
posed absence of simuliids although other biting flies such as CulcZcoidesJ,
Stomorys, Di~achlorus and mosquitoes were present.
Control of leucocytozoonosis can be achieved by locating :urkey
flocks at least 12.8 km (8 mi) from known breeding locations of potential
vectors and in areas free from possible wild turkey disease reservoirs.
Control of black flies by large scale aerial treatment of streams with
larvicides has been shown to markedly decrease the level of
parasitemias of L. emithi in sentinel turkeys (Kissam et al., 1975).
Fine mesh wire screen enclosures might be a valuable preventive tech-
nique with smaller flocks. Clopidol, a coccidiostat, at .025 and
.0125% in feed is effective in reducing the number of L. smithi para-
sites in turkeys as indicated by blood smears (Siccardi et al., 1974).
Investigators of leucocytozoonosis in turkeys in Florida have
studied the presence, consequence, and distribution of L. smithi in
the State but have not proven specific vectors. Travis (1939) reported
that nearly 100% of the wild turkeys in low areas of Georgia and Florida
were infected with L. smithi and suggested that an aquatic breeding
insect might be involved. Simpson et al. (1956) investigating out-
breaks of L. smith in turkey flocks near Palatka, Florida, reported
finding S. slossonae and a species #158 (=S. jonesi Stone and Snoddy,
1969) breeding near the turkey flocks and suggested that S. slossonae
might be an important vector. Davis et al. (1957) reported S. slossonae
as prevalent in Florida. Forrester et al. (1974) found a 72-75% in-
cidence of L. smith in 484 turkeys more than one month old sampled
from 10 localities in Florida. These authors also noted that a drop in
the infection rate of L. smithzi observed in wild turkeys in the Sikes
Fisheating Creek Wildlife Management Area and other areas of the State
in 1971 corresponded with low rainfall and low stream conditions and
theorized that reduced numbers of potential black fly vectors may have
resulted in the reduced prevalence of Leucocytozoon in turkeys.
MATERIALS AND METHODS
Black Fly Survey
Primarily, efforts in this survey concentrated on collecting larvae
and pupae from Florida's streams and rivers. In Alachua County ten
streams were selected and in nearby counties an additional twenty flows
were chosen which were fairly permanent and represented a diversity of
ecological conditions. An attempt was made to secure samples from each
of these streams at least once a month. Approximately the same location
was revisited each time to obtain some consistency in stream and sub-
strate conditions in order to judge seasonal replacement phenomena and
normal population changes. Other sites in south, southwest and west
Florida were chosen, generally along highways or country roads for ac-
cessibility, which were visited at least quarterly to obtain records at
all seasons of the year.
At each location specimens were removed with forceps or placed
still attached to small portions of grass, twigs and so on into four-dram
lip vials with neoprene stoppers. Each vial contained 80% ethyl alcohol
and a code number penciled on a small slip of paper. The immatures,
collected in 30 to 45 minutes were deemed sufficient to indicate numbers
and species present. Dark pupae, indicating an advanced state of develop-
ment, were placed in four-dram screw cap vials on a strip of paper toweling
moistened in the stream and were held alive in order to allow the adults
to emerge. Images could be reared very successfully in this manner from
older and frequently, even younger, more pale pupae. The forceps used
were secured to my wrist by a string to prevent their loss when my
balance was upset by the current, when I was wading and saw an occasional
water moccasin, and at other critical times. A 1.53 m long rake and a
modified, extendible, golf ball retriever were used to obtain substrate
samples and specimens where it was impossible to wade.
At each collection location stream dimensions, velocity, pH, tem-
perature, color, and substrate were recorded an special data cards along
with indications of the attachment substrates of the immatures and an
estimate of the population size. Temperatures were obtained with a
metal-cased pocket thermometer, range -lo to 490C (300 to 1200F). Initial
attempts were made to determine the pH of the streams using two portable,
electronic pH meters. After obtaining erratic, inconsistent results in
the field with the meters, pH papers (pHydrion papers Micro Essential
Labs, New York) were tried with more success. These pH papers come in
ranges such as 3.5-5.5 and 6.0-8.0 between I and 12 pH units with illus-
trated color differences at .5 pH unit intervals and are rated accurate
to .25 pH unit in buffered solutions. The papers were checked regularly
for accuracy with pH 4.0, 6.5, 7.0 and 10.0 buffered solutions. Very
acidic readings were confirmed by bringing chilled water samples into
the lab and checking them on a Beckman pH meter. Experiments with
electronic, propelleredGurley flow meters proved them too cumbersome to
set up and difficult to use in the streams which were often shallow and
clogged with submerged vegetation. The floating cork technique suggested
by Dalmat (1955) and others was used. A weighted cork 3.8 cm in diameter
which floated low in the water and was connected to a 3.05 m (10 ft)
long string was timed with a stop watch to determine the stream velocity
at different points where larvae and pupae were located. Most immature
black flies were encountered on trailing vegetation at or near the sur-
face and thus the floating cork velocity figures should reasonably
reflect immature habitat conditions. A ruler one meter long with
centimeter markings was used to determine stream depths and widths. The
widths of larger flows were estimated by pacing across them, when possi-
ble, or by pacing across a nearby bridge. Other omnipresent items on
collecting trips included an insect net an-d a snake bite kit.
Alcohol specimens and pupal vials from each site were placed with
the data card into cloth bags and transported to the lab where identi-
fications were made with the aid of a Bausch and Lomb stereamicroscope.
Larvae were usually identifiable without dissection (except unraveling
of the respiratory histablast); however representatives of each species
were dissected and mandibles, respiratory histoblasts, cephalic apotome,
gular notch, antennae and fans, and anal sclerite and hooks were pre-
pared in cellosolve and mounted in balsam under individual small cover
slips. Adults which emerged were allowed to harden for a few hours and
were then killed and placed in alcohol, prepared and mounted in balsam
on slides, or pinned on minutin nadeln inserted in white polyporous
pieces on #3 insect pins.
In addition to immatures, some adult collecting was undertaken
using a variety of traps. Black fly adults have been obtained in Florida
by other individuals using light traps and Malaise (flight) traps. In
this research three types of adult fly traps were used: the Manito'oa
trap (Thorsteinson et al., 1965), the blackout box trap (Anderson and
Dicke, 1960), and a modified ramp-type trap. The latter two types of
traps will be discussed with the Leucocytozoon. work. A modified Manitoba
trap (Fig. 1) was constructed using 3 bamboo poles, each 2.1 m (7 ft)
long, with a hole drilled vertically into the uppermost node of each
pole into which a leg of a metal tripod laboratory ring stand was in-
serted. A triangular serex or organza canopy 1.2 m (4 ft) tall was
positioned inside the leg frame and anchored at the three lower corners
by strings tied through a hole in each pole. The apex of the tapering
canopy with a 5.1 cm (2 in) wide elastic opening was drawn up a short
distance through the ring opening and stretched around the lower lip of
a 17.8 cm (7 in) tall lantern jar, the collecting vessel, which rested
on the ring. Th-e jar was topped with a double layer of white serex or
organza held in place with rubber bands. The jar contained an inverted
plastic funnel affixed to the jar with hot plastic glue. On strings
attached around the base of the jar and hanging into the canopy was
suspended a 1.2 m (4 ft) long cord with a black, cylindrical (20 cm
wide, 22 cm tall) metal can or a black cardboard triangle, 63 cm on
each side, on the end. A 23 cm wide black plastic "skirt" was sometimes
fitted over the top of the canopy and positioned at the lower margin of
the canopy to accentuate the "window" effect. A 31 cm long cloth bag
which contained 1.36-1.8 kg of dry ice was also hung inside the canopy.
The dry ice and the black target attracted Simuliidae and other Diptera
which flew or walked up the canopy into the collecting jar. Adults
could be aspirated from the jar by removing the outer cloth layer and
inserting an aspirator tube through a small slit in the lower layer of
material. Specimens could also be taken to the lab in the glass jar and
immobilized by cold in a refrigerator, though condensation inside the
jar often wet the specimens. Manitoba. trapping was accomplished using
one or two traps at a time in six counties at twelve locations for one
Figure 1. A modified Manitoba trap with a black plastic skirt.
to eight hour periods during the months of January through October. The
air temperature, wind speed and relative humidity were measured during
the collecting periods. Trapped adults were pinned or mounted on
slides as with reared adults. Identification of irmmatures and adults
was possible using taxonomic keys in Stone and Snoddy (1969), Stone
(1964) and smaller publications on individual species.
Leucoc~ytozoon amithi Transmission
In the investigation of the transmission of L. smith to turkeys
in Florida, modifications of Koch's postulates were used as guidelines.
The first aim was to establish an association in nature between the
host, the disease, and the vector. It was necessary to have a sus-
pected vector, preferably a clean, reared one, feed on an infected host
in the lab and to observe the stages of the parasite, especially the
infective stage, the sporozoite, in the fly. A clean host was then to
'be infected by a vector which had become infected in the lab with the
Leucocytozoon parasite being recovered in the gametocyte stage from the
formerly uninfected turkey.
In order to confirm the reported presence of L. smithi in certain
areas such as the Lochloosa Creek and Fisheating Creek Wildlife Manage-
ment areas (L. Williams, 1974 personal communication; Forrester et al.,
1974), I set out sentinel turkeys. The same was done to establish the
presence of the disease in other areas, and to obtain infected hosts to
serve as donors for transmission studies. These turkeys, like all clean
poults used in the transmission studies, were either hatched from fertile
eggs in a fly-proof room or obtained as disease-free one-day old poults
and held in a fly-proof room until use. Difficulties in obtaining clean
poults for experiments early in 1975 when winter and early spring species,
suspected to be L. smithi vectors, were flying were overcome for work
in early 1976. Usually three birds were placed out in the field near
a black fly stream in a cage for about a week and provided with food
and water (Fig. 2). There were three primary sentinel sites, all in
Alachua Co., and additional birds were also set out for short periods
at Fisheating Creek, Glades Co. (see Table 5). Sentinels were also
placed out on a few occasions in a ramp trap described below. In addi-
tion I worked with Dr. D.F. Forrester at Fisheating Creek during 1974
when he exposed many young turkeys for two week periods in cages in
cypress trees and on the ground and in 1975 when cages and ramp traps
were used for turkey exposures.
To determine the species of black flies generally present at the
sentinel sites, immature stages were collected and adults on the wing
were sampled with Manitoba traps as described in the previous section.
To identify black flies specifically attracted to turkeys I used blackout
box traps (Fig. 3), ramp traps (Fig. 4) and exposed turkeys (Fig. 5).
Anderson and DeFoliart (1961) used the technique of blackout box trapping
in Wisconsin. In Florida the blackout box traps were used at two
locations in 1974 and an additional seven sites during 1975 (see Table 7).
A turkey was exposed in a cage on the ground or on a platform in a tree
for about 15 minutes. A large cardboard box with two plastic collecting
cups on the top which appeared as bright areas inside the otherwise dark
box was then placed over the cage and bird. Black flies completing their
blood meals or unfed but on the turkey exhibited a positive phototactic
response, flew to the collecting cups and were captured. The flies were
removed to holding cartons, the turkey was re-exposed, and the process
;.j13 '' ~k9- );~x~i~
Figure 2. Sentinel turkeys in an exposure cage.
Figure 3. A blackout box trap in the field.
Figure 4. One view of a ramp trap.
Figure 5. An exposed turkey in the field.
t I (1~
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
elsewhere were quickly aspirated before they could scurry beneath the
feathers and were placed in the glass aspirator tube against the bare
skin of the underside of the wing elbow, the bare skin of the chest
over the rib cage or against the turkey's neck and allowed to feed to
repletion. The fly was observed, the feeding timed, and the turkey
soothed and kept as still as possible to assure a complete blood meal.
Once fed, the flies were transferred to paper pint ice cream cartons with
netting tops, provided a moistened paper towel to maintain high humidity
and transported to the lab. The exposed turkey technique was used to
collect black flies on numerous occasions from January through August
at 8 locations in 3 counties (Table 8).
To obtain black flies that had not been exposed to infections in
the field and to obtain specimens of known identity, especially when
working with species in the Simulium subgenus Phosterodoros which are
difficult to distinguish as adults and are much easier to separate as
pupae, adults were reared in the lab. Some adults were reared in pupal
vials from field collected pupae and other adults were reared form
field collected larvae and pupae in an aquarium with aerated water and
a fine mesh netting cover.
Black flies which fed on exposed infected hosts in the field were
held in the lab for at least two days after feeding and then allowed to
bite an uninfected bird. All adult flies maintained in the lab were
held in paper pint-sized ice cream cartons covered with netting and
provided with moist cotton for water and a raisin for nutrition. Up to
four cartons were kept at one time together in either a large plastic-
covered aquarium or a glass desiccator-type container (Fig. 6) where
high humidity was maintained by including a sponge half immersed in a
Figure 6. Glass container and paper cartons used for holding black
fly adults alive in the laboratory.
cup of water. Flies captured in an unfed condition in the field or
reared in the lab were allowed to feed on an infected turkey, held for
two to seven days and then allowed to feed on a clean bird. All
laboratory feedings involving uninfected hosts were conducted in a fly-
proofed former bull room at the Veterinary Science area at the University
of Plorida. This room was incandescent lighted,had a double door entrance
and had windows covered with very fine mesh organza material. Feedings
on infected hosts were conducted either in the Veterinary Science room
or at the fluorescent-lighted Veterinary Entomology Lab on campus.
Black flies were routinely taken off their sugar source at least 24
hours before planned feeding trials and maintained only on the water
provided by the moist cotton. Black flies were aspirated from the car-
tons or chilled in a refrigerator and removed with forceps and placed,
usually ane per vial, into seven-dram, clear plastic, cylindrical vials
covered with green, fine mesh (30/6.45 sq cm) netting at both ends and
held against the bare skin of the imrmobilized turkey. Room temperature,
relative humidity and feeding times were recorded.
Uninfected turkeys bitten by possible vectors were held in a fly-
proof room and the appearance of gametocytes in the peripheral blood
was monitored by blood smears obtained by wing vein punctures every 3
to 4 days. Blood smears were processed by air drying, fixing in
absolute methanol and staining in a 1:50 dilution of Giemsa solution
and distilled water. Flies were dissected in .9% physiological saline
and midguts and salivary glands were observed for the presence of
oocysts and sporozoites of L. smith. A few midguts were fixed in 10%
formalin, stained in Haematoxylin and mounted in balsam, with the
necessary intermediate steps for dehydration and clearing, in attempts
to more readily view the oocysts. Most midguts were air dried, fixed
in methanol and stained with Giemsa, as were the salivary glands. The
preparations were covered with permount and a cover slip for microscopic
examination. Other fly structures were placed in 10% Na0H (except the
wings) for 48 hours, rinsed in water and acid alcohol, placed in 100%
cellosolve for 15 to 30 minutes and mounted in balsam on a microscope
RESULTS AND DISCUSSION
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
Table 1. Florida black fly species.
Onephia (Chephia) ornithophilia Davies, Peterson, and Wood*
SimuZium (Byssodon) meridionate Riley
SimuZium (Byssodon) slossonae Dyar and Shannon
SimuZium (Eusimuiwnw congareenamrw(Dyar and Shannon)
SimuZium (Phosterodoros) diziense Stone and Snoddy*
SimuZium (Phosterodor~os) haysi Stone and Snoddy*
Sinulium (Ph~osterodoros) jenningsi Malloch
Simulium (Phosterodoros) jonesi Stone and Snoddy
Simutiurm (Phosterodoros) Zakei Snoddy*
Simulium (Phosterodoros) notiate Stone and Snoddy*
S~imultiwn (Phosterodoros) nyssa Stone and Snoddy
SiimuZium (Phosterodoros) tacodium Snoddy and Beshear*
SimuZiwn (Psitozia) vittatum Zetterstedt*
Simutium (Simuliwn) decorum Walker
SimuZiwon (Simutium) tuberosum (Lundstriim)
SimuZiwn (Simulium) verecundum Stone and Jamnback*
Onephia species undetermined No. l*
Simutium species undetermined No. 1*
New collection record for Florida.
Figure 7. Locations in Florida where black flies have been collected.
Table 2. Florida black fly distribution records by county.
Alacua 3 x xx x x xx x x
Baker~Q 1 xx
Dade 1 x
x x x xx x
x x x
x x x
x x xx
x x xX xx
x x X
x x xX x xx
xX x xx
Pinlla 1\ x
Sute 1 x x
Waul 1 xO~rO 3 ~
Wralng 4 x x xx
Table 2. (Continued)
By reading across or down from a selected species the number of times
the immatures of that species were collected together with the immatures
of another species is indicated by the figure at the intersection of the
Table 3. Black fly associations based on collections of immature stages.*
26 1 4 4
84 3 1 38107 73
2 19 7
2 19 92
19 24 5 19 36
S. slpp. undetermined
2 2 32 10
52 15 186 55
6 2 49 23
73 4 21 8
2 1 40 11 3 253 44
150 15 54 19
8 44 10
8 9 50 33
4 50 6 78
52 6 73 40 150
15 2 4 11 15
186 49 21 5 21 253 54
55 23 4 8 44 19
s s ~o
Table 4. Black flies captured in Manitoba traps.
1974: 3 May
1975: 24 Jan
Univ. of Florida,
Preserve, Site 24
Preserve, Site 24
Junction SR 225/
340, Site 8
Preserve, Site 24
Creek, Site 1
Univ. of Florida,
Preserve, Site 24
Univ. of Florida,
Table 4. (Continued)
1975: 26 Jan
Double Run Creek,
8 km west of
Highway 115C on
Rt 90, Site 72
Double Run Creek,
Junction SR 225/
340, Site 8
Preserve, Site 24
Double Run Creek,
Date Location times (all females)
1976: 24 Mar Turkey Creek, 1640-1830 S. congareenzarum,
Site 216 S. slossonae
26 Mar Sante Fe College, 1030-1800 S. tuberosum,
Site 28 S. slossonae
Table 4. (Continued)
C. (C.)I ornithophilia
S. (B. ) meridionate
S. (B.)I slossonae
S. (E.)i congareenarum
S. (P.)I didiense
S. (P.)I haysi
S. (P.) jenningsi
S. (P.) jonesi
S. (P.) Lakei
S. (P.) notiate
S. (P.) nssa
S. (P. ) taxodium
S. (P.) vittatum
S. (.S.)I decorum
S. (S.) tuberocum1
S. (S.)I verecundum
C. spp. No. 1
S. spp. No. 1
A S' O N D
J F M A M J J
Figure 8. Seasonal occurrence of black flies in Florida.
with an indication of the species found in each county. Table 3 pre-
sents information on the frequency of association of black flies in
Florida's streams and rivers. Figure 8 shows the seasonal occurrence
patterns of the collected Simuliidae. Table 4 presents information on
species captured in Manitoba traps. Further reference to these figures
and tables will be made in discussing the individual species below.
Keys to the black flies of Florida, in all stages, are included.
Introduction to the Black Fly Keys
Characters which are important taxonomically on black fly larvae
and are noted on Figures 9, 10, and 11 include: the anterior, multi-
ple-rayed cephalic fans and their stalks (CF,CFS); the antennae (A); the
central dorsal section of the larval head capsule referred to as the
fronroclypeus or the cephalic apotome (CA) which bears usually dark
pigmented areas called head spots (HS); the ventral, setae bearing and
anteriorly toothed hypostomium or submentum (S); posterior to the sub-
mentum is the throat (or postgenal) cleft (TC) or gular notch which
differs in size and shape in the different species; laterally on the
thoracic region are a pair of organs called the respiratory histablasts (RH)
consisting of a number of coiled filaments whose number and branching
pattern are often diagnostic; the ventral proleg (VP); dorsally at the
posterior end the anal gills (AG), asmoregulatory organs, protrude and
may be simple and branlchless stalks or may have many branches and be
arborescent; behind the anal gills is located the anal sclerite (AS) or
X-piece which assists the larva in releasing its posterior from the silk
attachment patches; th7e cir-lfe or many rows of tiny anal hooks (AH) at
CF r A
Figure 9. Dorsum of the head capsule of a black fly larva (S. slossonae).
Figure 10. Venter of the head capsule of a black fly larva
(C. ornithophi. iia) .
Figure 12. Black fly pupa and cocoon (S. dixiense).
Figure 11. Lateral view of two black fly larvae (S. dixiense).
the very end of the larva; and the ventral protrusions which occur on
some larvae and are called ventral or anal tubercles (VT).
Characters important on the pupa include: the filamented repira-
tory organs (RO); tiny setae-like structures on the thorax called
trichomes and posterior dorsal tail hooks (Fig. 12). On the
cocoon lateral apertures (LA) or windows occur anteriorly in the
Phosterodcros species and the texture and general regularity of the
cocoon is sometimes useful in separating species.
On the adult wing the presence or absence of hairs under the sub-
costal vein (SC); hairs on the dorsal, basal portion of the radius (R);
the presence or absence of a basal cell (BC); and color of hairs on the
stem vein (SV) are valuable characters (Fig. 13).
On the adult female head the shininess or pollinosity of the frons
(F) and its shape and that of the clypeus (C), the color of the antennal
segments (A), and the shape or size of the sensory vescicle of the
maxillary palps (SV) are useful taxonomically (Fig. 14).
Important characters of the male genitalia include: the appearance
of the ventral plate (VP) and the shape of its median portion; the shape
and relative lengths of the basimere (B) and distimere or clasper (D);
the presence or absence of a basal lobe on the distimere; and the number of
distal spines (DS) or teeth on the distimere (Fig. 15).
On the female hind leg the presence or absence of a dorsal groove
called the pedisulcus (P) on the second tarsal segment and the size of
the calcipala (CL), a flattened lobe on the inner side and at the apex
of the basitarsad segment, as well as presence or absence of a basal tooth
(BT) on the tarsal claws are important characters (Fig. 16j).
Structures useful for determinations in the region of the female
Figure 13. A wing of the black fly Cnephia ornishophilia.
Figure 14. A frontal view of the head of a female black fly (S, notiaele.
Figure 15. The male genitalia of a black fly, onephia omithophilia.
Figure 16. The distal portion of the hind leg of a S. meridionatle
Figure 17. The terminalia of a female S. meridionate.
genitalia (Fig. 17) are the size and shape of the genital. fork stem
(CF~S) an1d arms (CFA), the ovipositor lobes, each anal lobe (AZL) and
eachi cercus (CR).
A key to ther larvae of the blacki flies of: Florida.
(Photographiic illustrations of certain structures referred to in this
key and the keys which follow are included in the sections on the in-
dividual species. The larval key is primarily of value for later in-
stars. For further illustrations; refer to Stone and Jamnlhackk (1955),
Davies e~t al. (1962), Dood et al. (1963), StonE (1964), Stone and
Snoddy (1969~), Snoddy and Beshecar (1968) and Snotddy (11971, 1.976).
la. Hypostol:iuim convex, along anterior margin; head sp~ot pattern as in
Fig. 18. . .. ..... .. .. .. .. . C. ornrithop~hilia
16. Hypostomiumum cone~a\ or leve~l alo~ng tol anterio~r maergin~; the hcad
spot pattern not as in Fig. 18 ... .. .. ... . ... 2
2a. (Ib) Hiead spots light (white) .. .. .. .. . ... . 3
2b. Nlead spots dark or indistinct in a fulv~ous pattern . ... .. 5
3a. (2a) He~ad s5pot:, conpsristing of a cenitral.1 poste-io~r w~hte spot1 with
dark rays projecting and dive~rginge anterio~rly (Fig. 247, Stone and
Snoddy, 1.969; nol mature larva!e coll~ected~). . ... .S. mier~c~'idone
3b. Hea:d spot! s with! 3nterjior anld po~ster~ior- medial anld lalteral spots all
whiLe, spots vi-ib~le in thie usual pofitjions . .. .. .. .. 4
4a. (3b,) Meiccll an~terior and postecrior wh~ite hanld spots bor-dered by a
darki fulvous area on each sidle (F'ig. 99); Iespiratory his8tchlost
wilhl 8 fijlownts~n .. . ,. .. .. . . . ,'. decoruim
Ab. Nol dist inc. dairki fulvous border b~y rcontra? l he spots; histoblnst
wJith 6 filamints; .... . .. . . . . S. :'''^wrecundu
5a. (2b) Gular notch in the-form of a shallow inverted -v; headlspots
as in F~ig. 121 ... .. .. .. ... . .. C. species No. I
5b. Gular notch rectanngular, sub-rcctangular or sagiittiform. . .. 6
6a. (Sb) Cul~ar nlotch broadly sagittiform, ext~eundin at least across 8
of the venter of the head capsule, as widle as long .. .. .. 7
6b. Cular notch not Ragittiformi or else longer than wide .. .. .13
7a. (6a) R~espiracory histabla~sts consisting of 10 filomntins 9 of which
arise fr~om a thick basal 10th (Fig. 60); aaterior medial head spots
often weak . .. ... .. .. ... .. .. S. Jonesi
7b. Respiratory histablast not as above; a.ll head spots usually dark,
distinct .. .. .. ... .. .. .. .. .. .. ... 8
8a. (7b) Respiratory histroblast with 6 filaments; anal tubercles not
prominent. .. .. .. .. .. .. ... .. . .. S. notiate
8b. Respiratory histoblast within more than 6 filarmen~ts; anal tubercles
promninenlt........ ... ... .... ........9
9a. (8b) Respiratory histoblast wait-h 7 filanones. . . .. S. haysi
9b. Respiratory histablast with more than 7 filame-ncs. .. .. .10
10a. (9b) Respiratory hristoblast with, 8 filaments. . ... S. tiaOicodiim
10b. Respiratory histohlast with more than 8 filamnents. . ... .11
11a. (10b) Rlespiratory histablast ulith 9 ;ilanments .. .. .. ,S. Zakei
11~b. Respiratorq histohlzst wJith 10 finments . .. .. .. .12
1.2a. (1lb) 10 recspira::ory h-iotablast filamennts w~ith a pattern as in
Fig. 81.; a1nal tuberele~s piromuinenl 91nd CocaiI. .. . S. niYouC
1.2b. 10 respiratory hiistchlast filametnts w~ith a palttorn as in Fic. 53;
anal tuibercler sma1 :l and ronded lc.. . .. . .. jennringsjri
13a. (6b) Gular notch long extending b the distance or more to thle teeth
of the submentum, sagittiform or subrectangular, pointed or rounded
anteriorly. .. ... .. ... .. ... . . . 14
13b. Gular notch extending less than k the distance to the teeth of sub-
mentum, subrectangular. ....... ...... .. 16
14a. (13a) H-ead spots indistinct in fulvous area; gular notch usually
with parallel sides; anal tubercles inconspicuous . S. tuberosure
14b. Head spots distinct; gular notch with curved margins, elongate
sagittiform; anal tubercles prominent . . . . . . 15
15a. (14b) Re~spiratory histablast with 6 filaments; larvae reddish.
. . . . . * S. slossonae
15b. Respiratory histoblast with 10 filamren~ts; larvae ye~llow.
. . . . . . . . . . - S, clixience
16a. (13b) Respiratory histoblast filaments 4 in number; gular notch
longer than wide. . .. ... . .. . . S. species No. 1
16b. Respiratory histoblast. filaments numbeLr more than 4; the gular
notch not longer than wide. .. .. .. .. ... . .. .. 17
17a. (16b) Respiratory histablast with 12 filaments; the anal tubercles
are large, prominent; a medium~-sized larva, 6 mm~ long.
. . . S. conga~ailreeam
17b. Respiratory histoblast withi 16 filaments; the anal tubercles are
absent or inconspicuous; a large larva, 8-9 mml long . S. viCttatumi
A key to the pulPae of the black flies of Florida.
(No pupae of Cnephiia species No. 1 have been observed.)
la. Cocoon a loose mass of silk, indistinct shape; strong dorsal books
at the posterior end of the pupa. . . .. .. C. ornithophilia
lb. Cocoon a distinct slipper or pocket shape, strong dorsal hooks
absent. .. .. .. .. .. .. .. .. .; . . .. . 2
2a. (lb) Cocoon with large anterior, lateral apertures .
2b. Cocoon uniform without large lateral apertures. ...
3a. (2a) Pup~a wath 6 respiratory filaments on each side;
antero-ventrally comlpletely joined (FiG. 75). ....
3b. Pupa with miore than 6 filaments ...........
4a. (3b) Pupa with 6 filaments rising off a strong basal
(Fig. 48) . . . . . . . . . . .
Ab. Pupa with more tha7n 7 fil.aments ...........
Sa. (4b) Pupa with 8 filamecnts with four pairs of 2. ..
5b. Pupa wit-h more than 8 fi'l~anclts ..........
. . . 3
. . .10
. . .6
6a. (5b) Pupa with 9 filaments in the pattern 2, 2, 2 and 3 from the
dorsal. .. .. ... .. .. .. .. .. .. .. .. S. lakei
6b. Pup3 with 10 filaments. ... .. .. ... .. .. .. 7
7;1. (6b) Pupa wJithl 9 filaments rising from a thick basal 10h one
(Fig. 60) ... ... . .. ... . ... .. ionesi
7b. Pupal respiratory organ lacking a str-ong basal filnment . .. 8
88. (7b) Pupa w~itn 10 respiraltoryy filamecnts whichh rise as 5 pairs, the
petiole for filaments 7 nnd 8 (from; thIe dorcal) noticiably longer
and stouter than the other petioles (Fig. 12) .. .. S. diziense
8b. Pupa with 10 filaments, some of which~ rise in pairs others in
triplets. .. .. .. .. . .. . .. . .. 9
9a. (8b) The lower more ventral respiratory filaments on long petioles,
filaments long. .. ... ... .. .. .. .. .. S. nyssa
9b. Lower filaments rise close to the base, petioles short, filaments
short .. .. .. ... ... .. .. . S. jcinnngsi
10a. (2b) Pupa with 4 respiratory filaments. .. .. S. species No. 1
10b. Pupa with more than 4 filaments .. .. ... .. .. 11
11a. (10b) Pupae with 6 filaments in the respiratory organ1. .. 12
11b. Pupae with more than 6 filaments. . . ... . 14
12a. (11a) Pupal cocoon with concave anterior edge (in lateral view);
respiratory filaments rising from long petioles (Fig. 28)S. slossonae
12b. Pupal cocoon with convex or straight anterior edges; petioles
short ... .. .. .. ... . .. .. . ... .. .3
13a. (12b) Respicatory filaments widespread, base of filamlents 5 and
6 sharply separated from banse of 3 and 4,filaments long (Fig.115).
. . . . . . * * S. vercun dw~n
13b. Respiratory filaments not: widespread, bases about equidistant,
filaments short (Fig. 108). ... .. .. ... .. S. :ubcr'oswni
14a. (11h) Pupa with 8 respiratory f~laments; cocoon with rouGh text~ure.
. . . . . . . . . . S. accr:C r nl
14b. More than 8 filaments . .. .. . .. .. .. . 15
.15a. (141,) Pupa with 12 respiratory filaments; cocoon with long anterior-
dorsal projection (Fig. 35).. . .. ... .. .. cong'areenarurr
15b. Pupa with more than 12 filaments; cocoon lacking dorsal projec-
tion .. .. .. .. .. .. .. .. . .. . .. .. 6
16a. (15b) Pupa with 16 filaments in 8 pairs . .. .. .. S. vittatus
16b. Pup3 withr more than 20 filaments in pairs and in 3's .S. meridionale
A key to the adult male black flies of Florida.
(Adapted and modified from Stone and Enoddy, 1969)
la. Second tarsal. segment of the hind leg without a pedi~sllcus but a
shallow~ depression may be present; radius vein with hair~ dorsally
on the basal segment; large basal cell present . .C. ornithophilia
lb. Second tarsal sfement of the~ hind leg with a deep pedisulous; base
of radius withi or without hair dorsally; large basal cell absent.
2a, (Ib) Basal portion of radius with hair dorsally S. congareeniar~um
2b. Basal. portion of radius without hair dorsnlly. .. .. .. .. 3
3a. (2b) D~istimere~ short, stout, with; 3 or mo~re tceeth .. S. visttumw
3b. Distimore long, with 1 tooth or none .. .. ... .. .. 4
4a. (3b) Distimere with a rounded 10be on the inner margin near the
base .. . . .. .... .. ... .. ... .. 5
4h, Distimefre without a lobe or basal projection .. .. .. .. 6
Sa. (42) Distimuere with ba~sal lobe bearing a number of stout, spine-
Itke scenec . .. .... .. .. .. ... .. S. tuber'osur
5b. Distime~re with basal lobe bearing finec hairs only. . S. stossonae
6a. (4b) Ventral plate In ventral view broadly rounded. S. meridionale
6b. Ventral plate in ventral view more narrow, compressed from th~e
sides. . .. . ... . .. .. .. . .. .. 7
7a. (6b) Ventral plate with basal arms hearing distinct lateral pro-
jactions; posterior third of sc~utum shiny with indistinct hairs. 8
7b. Ventral plate with basal arms without distinct lateral projections;
posterior quarter or less of scutum shiny with some strong erect
hairs. ... .. .. ... . .. ... .. .. . .. 11
8a. (7n) Ventral plate in ventral view with median portion longer than
wide and parallel-sided or nearly so; scutumn with silver spots
narrow, oblique; thle dark area be~tween the spots broadens anteriorly.
. . . . . . . . . . . . . . S. dizience
8b. Ventral plate in ventral view with median portion not longer than
wide and' widened toward the end; anterior silver spots ;nd inter-
vening dark area variabLe. .. .. ... .. .. ... .. 9
9a. (8b) Scutuml with large anterior silvery spotsa each extendingi about
one-third the distance across the front of thle scutum; thle dark area
between the spots na1rrows strongly to thle anterior margin. .S. Jone~d
9b. Scutumi with smanller silvery spots; ilhe dark area betweenr th:e spots
is broad convergies little, and is narrow~est a-bout mindway along
the l~ength of the spot; .. .. .. .. ... . S. hiaUGL
10a. (7b) Ventral plate in ventral view narrow, V-shaped, middle
region tapers almost to a1 point; with a1 ventral keel. S. decorumr
10b. Ventral plate broader, not as strongly compressed in the middle
section; ventral keel absent. .. .. .. .. .. S. vereoundlun
A key to the adult female black flies of Florida.
(Adapted and modified from Stone and Snoddy, 1969)
la. The second hind tarsal segment lacks a pedisulcus, although there
may be a slight depression; the basal cell is present and the basal
portion of the radius bears setae dorsally . .. C. ornlithophiliai
lb. The second hinid tarsal segment with a deep pedisuicuis; large basal
cell absent and the basal portion of the radius bears or lacks setae
dorsally (Simu~7~ium.. ... ... .... . . . . ..
2a. (lb) Radius vein with hair dorsal~ly on the basal section.
. . . .. . . . .. S congareelnarumn
2b. Radius without hair dorsally on the babal section. .. .. .. 3
3a. (2b) Tcrsal claw with a prominent basal. tGOoc .. .. .. .. 4
3b. Tarsal claw simple, without a prominent basal tooth. .. .. 5
4a. (3a) Fr-ons gray pollinose .. ... .. .. .. S. mieridionale
Ab. Frons shiny black. .. .. ... .. .. ... .S. slossonna
Sa. (3b) Frons and terminal abdominal targites shiiny black or dark
brown. ... . .. .. . . .. ... . .. . .. 6
5b. Fron~s and tecrminal abdominal tergites at least lightly pollinose.10
6a. (Sa) Sub~cosr-al vein with a row of hai~rs ventral3y; scutumn
subshiny .. .. .. .. ... . . .. . .. .7
6b. Subcostal vein without hairs ventrally; scutumi shiny .. .. 8
7a. (6a) Fore tibia with a narrow gray-white patch not more than one
third the width of the tibia; inner margins of ovipositor lobes
fairly st-raigiht. .. .. .. .. .. .. .. ... S. tubhero~swn
7b. Fore tibia with a brilliant white patch which covers one half or
more of the tibia; inner margins of the ovipositor lobes concave
enclosing anr oval area .. .. ... ... .. S. ver~ecundum
8a. (6b) In anterior view the scutum displays a pair of fairly distinct
rounded pollinose spots with a darker area of the scutum in between;
hairs on the stem vein are dark brown to black . .. S. jenningsi
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
9b. Clypeus longer than wide . .. . .. .. .. S. dixiense
10a. (5b) Scutum silvery gray with dark brown markings; abdomen with a
black and light gray pattern .. .. .. .. .. S. v~ittatusm
10b. Scutum uniform brownish gray without contrasting dark brown manrkings;
abdomen black with thin g~ray pollinosity without a pattern.
. . . . . . . . . . S. decolmm
Introductionto th Individual Species Sections
The followingf sections deal specifically with the black fly species
found in Florida. References are listed below each species name which
provide synonyms for the currently accepted name and additional sources
of descriptions andi figures. The descriptions which follow the brief
taxonomiy portions are intended to point out outstanding or diagnostic
characters and are accomlpanied with photographic illustrations. Under
"Florida Observations" below the heading "Stream Parameters" A summary
is given of the dimension, pH, temperature, and velr.city figures for
streams which contained each par-ticular species. i~n the sectionn on
Florida collection records the numbers i:amediatel~y inl!.owing the county
names refer to individual collection siteS identified inl the A~ppendix.
The code for thle collection records is as Follow~s: S refers to small,
very young larvae with no or weakly developed respiratory histoblasts;
M refers to medium-aged larvae with distjinct but white histablasts;
Refers to large, mature larvae withi dar:k histobl~asts; P refers to
full pupae or pupal exuviae; C refers to ucocons; anld A1 refer~s to male
or female adults. Tlh- majority of records refer to specimens the
authonr personally collected; in those instances where specimens were
gathecred by other individuals, the collector is noted. On the indi-
vidual species ma~ps wJhen a record is included with the only locality
information beingi the county the mark for that record on the ma~p is
placed in thle center of the :ounty. Immat~nure, very young larvae
lacking well-develouped respirnatry h~istobtasts in the subgenus
Phol~sterodoros~? are difficult if not impossible to differentiate as: are
the adlul~t fema;les. Records for t~he P~hostr:: odo:. l~.i ronpeces are based~
prima~rly oin mature larvae and p~uPre. Only at locntionss whero
essentially a single Phosterodoros species was found to exist are
records for earlier stages included. Records for undetermined species
are listed as S. (Phosterodoros) spp. in the Manitoba trap results
(Table 4). Dr. E.L. Snoddy kindly examined immature and adult specimens
of S. jenningsi, S. lakei and S. tarodium~ from Florida and confirmed my
Onephia (Cnephia) or-nithophilia Davies, Peterson, and Wood
onephia ornitshophiziai Davies, Peterson and Wood, 1962, Proc. Entomol.
Soc. Ontario 92: 102 (female).
Onephzia ornithoph2iia Stone and Snoddy, 1969, Auburn Univ. Agr. Exp.
Sta. Bull. 390: 25 (female, larva).
Taxonomy. Davies et al. (1962) first described a female of this
species. The holotype was collected from a blue jay at Algonquin Park,
Ontario and was deposited in the Canadian National Museum. Paratypes
are located in the U.S. National Museum. Stone and Snoddy (1969) in-
dicated the male of C. orn)ithophlZ~ia was not known and that the larva
and pupa were apparently indistinguishable from those of C. pecuaruo.
Description. The larva is large, about 8 mm long with a grayish
brown abdomen and a light yellow brown head capsule. The cephalic
spots are dark with the median groups of about 20 forming one long
continuous row (Fig. 18). The submentum is convex apically and bears
small teeth. The gular notch has fairly parallel sides, is broadly
rounded anteriorly and extends less than 1/3 the distance to the sub-
mental teeth (Fig. 10). The cephalic fans each contain about 60 rays
with long hair-like spines. The anal tubercles are absent.
Figure 18. The head spots of a larva of C. ornithophilia.
Figure 19. The pupal exuvium and cocoon of C. ornithop~hilia.
The pupa bears strong dorsal posterior hooks and is located in an
irregular, loosely woven cocoon. The respiratory organs each contain
about 30 filaments (Fig. 19).
This is believed to be the first description of the male. The
wings are 3.75-4 mm long. There are hairs dorsally on the base of the
radius. The first segment of the flagellum is the longest, about twice
the length of the other segments. The scutum is dull brownish black with
numerous short golden hairs. The scutellum is reddish brown and bears
long dark hairs. The legs are almost uniform yellow brown. The second
hind tarsal segment lacks a pedisulcus. The abdomen is dull brownish
black dorsally and lighter gray or light brown ventrally. The ventral
plate in ventral view is broad and the tapering distimere which is
almost equal in length to the basimere has a single tooth at its apex
as in Fig. 15.
Plesiotype: Male, with associated pupal exuvium and cocoon,
NW 23rd Ave. and NW 83rd St., Gainesville, Alachua Co., Florida, 2
February 1975 (Pinkcovsky), to be deposited in the U.S. National Museum.
The wings of the female are about 4.5 mm long. The base of the
radius is covered with setae and a basal cell is present on the wing.
The frons bears short yellow hair. The pedisulcus is essentially absent
and the prominent tooth on the basal claw has convex margins (Fig. 20).
The apex of the abdomen is shiny. The genital fork has widespread
narrow arms as in Fig. 21.
Distribution. Stone and Snoddy (1969) indicate C. orni-thophiia
occurs in Louisiana, Mississippi, South Carolina, Virginia, and Ontario.
Tarshis and Stuht (1970) report this species from Mlaryland.
Life History. Stone and Snoddy (1969) mention that C'. ornith~ophilla
Figure 20. Tarsal claw of a female of C. ornithophilia.
Figure 21. Genital fork and terminalia of a
is probably univoltine. Eggs collected in stream bottom samples and
placed in laboratory rearing tanks yielded first instar larvae three
days later (Tarshis, 1973). Larvae have been collected between 11
January and 18 March in South Carolina and adult females were reared on
6 February from pupae; females have been collected from 6 February to
1 May in Louisiana and Mississippi (Stone and Snoddy, 1969). Tarshis
(1973) working in Maryland found larvae from 14 November to 29 April,
pupae from 11 January to 22 April and adults from 2 to 17 March. Tarshis
and Stuht (1970) captured adult C. ornithophitia between 28 February
and 1 April in emergence cages over a stream in Maryland. When first
to third instar larvae were collected and reared in the laboratory in
aerated water at 15-220C pupae appeared after 2 to 21 days and adults
emerged 4 to 23 days after introduction of the larvae (Tarshis, 1973).
Tarshis (1973) found adults emerged as soon as 2.5 hours after pupation
[very difficult to believe] and that the number of males emerging ex-
ceeded the number of females emerging only on the first day.
Ecology. Tarshis and Stuht (1970) found massive numbers of larvae
on leaves, rocks, and twigs in a stream that flowed .153 to 1.0 m/sec
(.49-3.3 ft/sec) from a pond through an oak and willow woods. TIhe
stream was .92-1.53 m (3-5 ft) wide, 2.5-20 cm (1-8 in) deep and had a
pH reaction between 6.9 and 7.4. In a similar stream that was slightly
deeper and 1.22-3.66 m (4-12 ft) wide, larvae were collected at water
temperatures of .5-15oC (Tarshis, 1973). Tarshis (1973) mentioned that
pupae easily suffered injury and that development to adults did not
occur when pupae were removed from their substrate. Larvae of C. muitaba,
P. gibsoni, S. decorum, S. tuberosumn, S. venustum, and S. vitt~atum were
collected with C. ornithophilia in a pond runoff stream where
C. ornithophilia predominated (Talrshis antd Stuh~t, 1970).
Habits. Tarshis and Stubt (1970) suggest that C. ornithophilia
ov-iposits on the stream surface and the eggs sink to the stream bed.
Bennett (1960) in Ontario recovered C. ornithophilia (as CnephiJa "U")
fromt jays, hawks, robins, sparrows and other woodland birds. Tarshis
(1972) fed C. onnithophiia on geese and ducklings in the laboratory
and transmitted L;. simondi to ducklings by the bite of the fly. Tarshis
(1973) did not collect any C. ornithophi~lia from ducklings exposed on
the banks of a stream containing larvae and pupae of the black fly and
suggests that thle target hosts were inappropriate or perhaps that the
flies were absent at ground level, and concentrated in the canopy.
Width Depth pH Temperature Velocity
Mleanl: 2.57m 23.64 cm 4.7 14.70C (58.60F) .45 m/sec (1.47 ft/sec)
Mlin: .3 1.3 3.6 8.3 (47) .23 (.75)
Max: 11 90 6.6 24.4 (76) 1 (3)
C'nephia ornithophilia-lc is reported here for the first timne from
Florida. Both Stone (19)65) and Stone end Snoddy (1969) mention that
C. pecuarum occurs in Florida. Key chiaracters, such ats the shape of
the tooth on the tarsal claw of the female, on specinens I obtained in
Florida differed from those of C. pccuarwn.m In the U.S. National
Museum I located one adult On~ephia specimen from Florida, listed for
Site 204 below,, which was a male that had been labeled as C. pecuarum
but C. ornithophiliadn was pe~nciled inl underneath.. Adult and immature
specimens whlichl I submitted fro~m five collections representingli two welst
Florida (Hol~mes and Liberty) and one east F~lorida (Alachua) counties werre
identified by G.E. Shewell of the Canadian Biosystematics Research
Institute as C. omnithophilia.
Onesphia ornithophilia was collected from 31 sites in 14 counties
in Florida (Fig. 22). Most of the records represent collections of only
a few, usually young larvae with small, pale histoblasts, but the larger
overall body size compared to other species and distinctive head spots
and gular notch are discernible even in young larvae. Larvae, pupae
and associated adults were obtained at three sites 28, 119 (Fig. 23),
and 147). This species was first collected in the streams on 28 October
and last collected on 18 April. Most collections occurred and the
largest populations were encountered during December through March.
Collections of C". ornithophilia at a number of sites were made during
one winter month and the species was not found again until about the
same time the following year. It appears that C. ornithophilia is
univoltine in Florida. This species occurs in cool, fairly shallow
flows, usually at most a few meters wide, most frequently with a velocity
of .305-.61 m/sec (1-2 ft/sec) and a pH which normally is below 5. Im-
matures were also collected at Lebanon Station (Site 139) and Yellow
Water Creek (Site 70) where the pH approaches neutrality. At most sites
there was considerable vegetation especially trailing grass in the stream
to which the immatures were attached. At Blues Creek (Site 2) and Site
28 sparce vegetation was present. Site 28 originated in a small swamp
just upstream, was about .5 m wide, 10 cm or less deep and tumbled down
a root-filled and rocky bed through woods. In January a heavy popula-
tion of C. ornithophilia covered rocks, dead tree leaves, pine needles,
and a few blades of trailing grass. Masses of 40 or more larvae en-
circled twigs about .6 cm in diameter in the current. Most of the