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Spiroplasma Diversity in Two Tabanid Flies and in vivo Interactions of Spiroplasma and Entomoplasma
LaTonya T. Brown-Derrick
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SPiROPLASMA DIVERSITY IN TWO. TABANID FLIES
AND In Vivo INTERACTIONS OF
Spiroplasma AND Entomoplasma
LaTonya T. Brown - Derrick
■ 'v..;- W, itf- )" (5 G^c't t fii*" 4J Z*ch S. H f pd^r?rn Jihresy 6) $ ^ P // Spiroplasma Diversity in Two Tabanid Flies and In Vivt
Interactions of Spiroplasma and Entomoplasma
by
LaTonya T. Brown-Derrick
B.A., Columbia College, 1996
A Thesis Submitted to the Faculty
of the College of Graduate Studies
at Georgia Southern University
in Partial Fulfillment of the
Requirements for the Degree of
Master of Science in Biology
Statesboro, Georgia
March, 1998 Spiroplasma Diversity in Two Tabanid Flies and In Vivo Interactioi
of Spiroplasma and Entomoplasma
by
LaTonya T. Brown-Derrick
s'/as'A/f kne Van Tassell ! Date Associate Vice President for Academic Affairs and Dean of Graduate Studies Acknowledgments
I would like to thank the Chairman of my Graduate Committee, Dr Frank
French, for all of his advice and support, I would also like to acknowledge my
Committee members, Drs. William Irby and Quentin Fang, for their valuable assistance
I would be remiss if 1 didn't thank the following: Becky Wilson for her laboratory assistance and companionship during the long days, Ruth Braddy for specimen collection, and Mrs. Lorraine French for the wonderful cookies!
Finally, I express my appreciation to my husband, F Maurice Derrick, and parents, Mr. and Mrs. Moses J. Brown, for their love and support during the past year and a half
i Abstract
Chrysops vittatus, a small deer fly, and Tabamts atratus, a horse fly, were captured concurrently in Bulloch County, Georgia during June through September of
1989-1997 Cultures were made from adult females, and the frequency and diversity of spiroplasma carriage were determined by deformation and endpoint testing.
Of the 90 Chrysops vittatus processed, 23 (25 .6%) carried spiroplasmas. The most prevalent spiroplasmas were representatives of group Vlll (20/23): strain B1357
(7/23); Spiroplasma syrphidicola (5/23); S. chrysopicola (4/23); strain TAAS-1 (4/23).
Groups XVIII and XXXV were represented by S. Htorale (2/23) and strain B2649
(1/23), respectively.
Of the 47 Tabanus atratus processed, 22 (46.8%) carried spiroplasmas. The most prevalent spiroplasma was represented by S. tabatudicola (12/22) of group
XXXIII Other isolations were as follows: group XXIII, S. gladiatoris (3/22), group
IV, strain PPS-1 (1/22); group XIV, S. corruscae (1/22); group XVIII, S. htorale
(1/22); group XXXIII, strain TABS-2 (1/22); group XXXIV, strain B1901 (1/22), group XXXV, strain B2649 (1/22)
Tabanus longiusculis and Tabanus hneola were live captured and randomly assigned to five treatment groups: A - spiroplasmas only; B - entomoplasmas only,
ii iii C - spiroplasmas first and entomoplasmas later, D - entomoplasmas first and spiroplasmas later, E - control (sucrose only) Strain TN-1, Spiroplasma htorale, and strain ELCN-1, Entomoplasma e/lychniae, were administered in 5% sugar water.
Spiroplasma litorale was recovered from the following treatments: A - 12 of 12 gut cultures and 1 of 12 hemolymph cultures, C - 9 of 9 gut cultures and 1 of 9 hemolymph cultures; D - 7 of 7 gut cultures and 1 of 7 hemolymph cultures Entomoplasma e/lychmae was recovered from the following treatments: C - 5 of 9 gut cultures and 1 of 9 hemolymph cultures, D - 7 of 7 gut cultures and 1 of 7 hemolymph cultures
To my knowledge, this is the first report of laboratory induced infections of A litorale and E ellychniae being administered in sugar water Also, the gut and hemolymph samples where both S. litorale and E. ellychniae were recovered represent the first reports of concurrent mollicute infections in tabanids Table of Contents
Acknowledgments i
Abstract ii
Introduction j
Materials and Methods 7
Test 1 Natural Carriage 7
Test 2 Laboratory Transmission 9
Results 12
Test 1 Natural Carriage 12
Test 2 Laboratory Transmission 13
Discussion 15
Test 1 Natural Carriage 15
Test 2 Laboratory Transmission 16
Literature Cited 19
Tables 29
Figure 32
Appendices 33
IV List of Tables
Table 1
Spiroplasmas isolated from the abdominal viscera of 29
Chrysops villains and Tabanus alralus captured in Bulloch
County, Georgia
Table 2.
Spiroplasmas isolated from Chrysops villains and 30
Tabanns atralns compared with data for Chrysops bnm/eyi.
Hy bom i Ira difficilis. and Ta banns lineola captured in Bulloch
County, Georgia.
Table 3.
Number of successful Spiroplasma htora/e (Sf) and 3 1
Tjitomoplasma ellychniae (Ee) isolations made from two
Tabanus spp for five treatment groups
v Figure
Figure 1.
Mollicute Isolations From Gut and Flemolymph
VI List of Appendices
Appendix 1
Isolation Procedures
Appendix 2
Preparation of R.2 Medium
Appendix 3
Serological Testing
Appendix 4
Six Different Combined Antisera for Screening
vn Introduction
Spiroplasmas are helical, wall-less, motile prokaryotes found in the division
Tenencutes, class Mollicutes, and order Entomoplasmatales When first recognized,
they were confused with both spirochetes and viruses because of their morphological
similarities (Tully and Whitcomb, 1991). Spiroplasmas have been found in association
with arthropods, especially insects and ticks, and are the infectious agents of several
plant and honey bee diseases. The genus Spiroplasma was established in 1973 by an
international research team studying a helical mycoplasma found in plants now known as citrus stubborn disease (Saglio et til., 1973)
Several species of spiroplasmas have been found in the phloem sieve-tubes of plants. Spiroplasma citri became the first mollicute of plant origin to be cultivated and characterized (Cole et al., 1973). Spiroplasma kunkelii, the causative agent of corn stunt disease, was cultured in 1975 from Zea mays (Indian corn) and fully characterized in 1986 (Saillard et al., 1984). Spiroplasmaphoeniceum was naturally cultured from infected periwinkle plants in Syria and described in 1986 (Saillard et al., 1987)
Insects are particularly abundant sources of spiroplasmas, and most species of spiroplasmas are insect-derived. Isolations have been made from the insect orders of
Coleoptera, Diptera, Hemiptera, Homoptera, Hymenoptera, and Lepidoptera. While most spiroplasmas appear to have a commensalistic relationship with their insect host,
1 2 some infections produce a fatal response, e.g., S. melliferum and S. apis are honey bee pathogens They migrate from the insect-gut barrier into the hemolymph, where they multiply and later kill the bee (Mouches et al., 1983; Clark et al., 1985) Spiroplasma apis has also been cultured from the surfaces of flowers growing near an affected beehive suggesting that spiroplasmas were deposited by infected bees (Bove, 1997).
Spiroplasma poulsomi was originally isolated from neotropical species of
Drosophila It is transmitted transovarially and kills the male progeny of infected females. Because of the sex ratio abnormality produced, S. poulsomi was given the name sex ratio agent or SRO (Poulson and Sakaguchi, 1961) Spiroplasmas also have been isolated from mosquitoes: S. sabaucliense from a pool ofAedes sticticus and A. vexans (Chastel et al., 1985, Albalain-Colloc et al., 1987), S. culicicola from the salt marsh mosquito Aedes so/licitans (Hung et al., 1987), and S. taiwanense from Culex tritaenoirhynchus (Clark et al., 1987, Albalain-Colloc et al., 1988).
Of the three known tick spiroplasmas, only S. minim, obtained from the rabbit tick, Haemaphysalis leponspa/ustns, has been found (experimentally) to be pathogenic to vertebrate animals such as chick embryos, new-born rodents, and adult rabbits
(Pickens et al., 1968). Spiroplasma strains SMCA and GT-48 were isolated from rabbit ticks in Georgia. Although they were found to be serologically identical, they induce different experimental pathogenicity. Strain SMCA produces high incidence of cataracts in new born rodents while strain GT-48 produces fatal encephalitis in rodents (Clark,
1964; Bove, 1997). 3 Tabanid flies (Diptera: Tabanidae) are rich reservoirs of spiroplasmas (Clark et ai, 1984, French et al., 1992) and are host to a wide variety of different mollicute
infections The system by which tabanids become naturally infected with spiroplasmas is
unknown. Most tabanid females require a bloodmeal for oviposition. Nevertheless,
both males and females consume water and carbohydrates daily for flight energy (Schutz
et a!., 1989; Tesky, 1990) It is believed that spiroplasmas are obtained at carbohydrate
sources through feeding on nectar, sap flows, and honeydew (aphid anal droplets)
(Wedincamp et ai, 1996).
There are 35 recognized groups of spiroplasmas associated with plants and
insects with 11 groups isolated from tabanids. Only three of the tabanid associated
spiroplasmas have been isolated from other insects, S. syrphidicola, isolated from a
syrphid fly, S. corrusca, isolated from a firefly beetle (Coleoptera: Lampyridae); S apis,
isolated from honey bees (French et al., 1992, Whitcomb et ai, 1997)
Nonhelical mollicutes such as Acholeoplasma and Entomoplasma have also been
isolated from both plants and insects. McCoy et al. (1979) and Eden-Green and Tully
(1979) cultivated both sterol-requiring and nonsterol-requiring nonhelical mollicutes
from flowers and the surfaces of various tropical plants. In a study by Clark et al.
(1986), the presence of acholeplasmas and other unclassified, nonhelical mollicutes in
insects was reported, thus establishing a link between nonhelical mollicutes on plant
surfaces and an insect reservoir. 4 Some of the first nonhelical mollicutes from plant surfaces were strain PPA from the "powder puff' plant, Calliandra haematocephala, and strains L1 and GF1 from the flowers of lemon and grapefruit trees (McCoy et ai, 1989). Eden-Green isolated
Acholeplasma axanthum and A. oculi from coconut palms infected with "yellowing disease" (Eden-Green et ai., 1979). These two nonhelical mollicutes had previously been isolated from cattle and goats only, yet their presence on plant surfaces suggested an insect association (Tully, 1989).
Entomoplasmas are small, nonhelical, nonmotile, pleomorphic, coccoid mollicutes. Entomoplasma ellychniae was the first nonhelical mollicute to be recovered from an insect. It was isolated from the hemolymph of the firefly beetle FJlychnia corrusca (Coleoptera: Lampyridae) (Tully et al., 1989; Tully et al, 1993). As previously stated, E. corrusca is also the only arthropod to harbor a spiroplasma known to be frequently associated with tabanids (EC-1 of group XIV) (French et at., 1992,
Whitcomb et al., 1997). Other entomoplasmas isolated from fireflies include E. luminosum and E. lucivorax isolated from the gut of adult Photinus margmalis and
Photinus pyralis and E. somnilux isolated from the pupal gut of Pyractonema angulala
(Williamson et al., 1990).
When fireflies and tabanids were cohabited, E. ellychniae was presumably transferred from naturally infected Ellychnia corrusca fireflies to tabanids at their common feeding site (Wedincamp, 1994, Wedincamp et ai, 1996) To date, there have been no reports of natural infections of E. ellychniae in tabanid guts and/or hemolymph 5 While the actual mode of infection for entomoplasmas and spiroplasmas is unknown, this suggests similar transfer in nature.
Spiroplasmas have been isolated from the crop, hemocoel, and salivary gland of arthropods (Clark el a/., 1984). Spiroplasma isolations from the gut and hemolymph of tabanids have been made with relative ease (Wedincamp el a/., 1996, Appendix 1)
There have been several serological procedures and diagnostic techniques applied to identify and classify mollicutes The serological tests include the growth inhibition (Gl) test, the deformation (DF) test, the metabolism inhibition (MI) test, and serologically specific electronic microscopy (SSEM) The diagnostic techniques include enzyme- liked immunosorbent assay (ELISA), immunoflorescent staining, and scanning electron microscopy (Chen el al., 1989; Williamson and Whitcomb, 1983) In selecting the appropriate test, cost efficiency and rapid results were desired as long as accuracy was not compromised.
The difference in natural infection assemblages among the species studied in detail indicate host specificity (French el al, 1996, Moulder el al., 1996; Moulder,
1997), that may be best referred to as "ecological" rather than "physiological" host specificity. The ecological specificity is most likely due to differences in carbohydrate sources of spiroplasma infections.
One objective of my research was to determine the frequency and diversity of spiroplasma carriage in Chrysops villains, a deer fly, and Tabanus at rat us. a horse fly
These two tabanid flies are commonly captured together during the summer months 6 (June - September) in Builoch County, Georgia. I wanted to determine if seasonality was an important determinant of spiroplasma infection in adult tabanids
Tabanids have shown no host specificity when mollicute infections were induced or transferred in the laboratory Spiroplasma montanense and S. corruscae (Railey and
French, 1990; French et «/., 1994, Wedincamp et a/., 1997) were successfully established as gut infections in more than a dozen species of Tabanidae (Chrysops spp..
Hybomitra spp., and Tabanus spp ).
Harper and French (1994) reported laboratory induced infection of.V montanense in the hemolymph of Hybomitra ciifficilis Attempts to repeat the experiments have failed (French unpublished data)
Another objective was to investigate the /// vivo interactions of spiroplasmas and entomoplasmas especially in the hemolymph of tabanids, I wanted to determine whether dual mollicute infections in the gut would promote the invasion of the hemolymph by one mollicute. Through these projects, I hoped to provide additional information on the acquisition, transmission, and residence of tabanid fly associated mollicutes Materials and Methods
All tabanids were captured live in Gressitt/Malaise traps and by aerial band- netting at four locations in Bulloch County, Georgia: Sullivan - 32° 22' 14" N, 81° 48'
04" W, Toole - 32° 30' 32" N, 81° 36' 33" W; Kennedy - 32° 3 1' 09" N, 81° 47' 57"
W, Franlorr - 32° 32' 3 1" N, 810 46' 07" W After capture, flies were placed in individual spiroplasma-free plastic containers, chilled, and transported to the laboratory
Test 1 Natural Carriage
To determine the diversity and frequency of spiroplasma carriage in 90
Chrysops villains and 47 Tabanus alratns, the flies were surface sterilized in 20 mL of
0.5% NaOCl containing two drops of Photo-Flo 200® and rinsed with deionized FbO
The last two abdominal segments were snipped and the abdominal viscera were removed using fine-tipped forceps. For flies captured between 1989-1996, the viscera were minced in M1D medium (Whitcomb, 1983) For flies captured in 1997, the viscera were minced in R2 medium (Moulder, 1997, Appendix 2). After mincing, the broth and tissue fragments were syringe filtered through 0.45 pm pores into a 3 mL plastic snap cap culture tube and incubated at 30oC
7 8 Upon acidification of the medium as indicated by a change in color from red to yellow, cultures were viewed by darkfield microscopy for spiroplasma cells. When a culture reached the logarithmic growth phase and demonstrated good morphology, it was subcultured and either held at -70oC or immediately analyzed by deformation (DF) testing (Williamson et ai, 1978, Whitcomb & Hackett, 1996, Appendix 3). For preliminary screening, each culture was tested against six different combined antisera
(Appendix 4). In individual wells of a microtiter plate, 20 pL of culture and 20 pL of combined antisera were mixed and allowed to react for a minimum of 30 minutes
Slides were made of the reaction mixture and viewed under darkfield microscopy at
lOOOx to determine whether the antisera caused morphological deformation or agglutination of spiroplasmas.
To determine the species of spiroplasma, an endpoint test was performed. For each combined antisera with which the spiroplasma reacted, 20 pL of culture and 20 pL of serial dilutions of the individual antiserum components ranging from 1:10 to
1:2560 were mixed and allowed to react for a minimum of 30 minutes The greatest dilution to have at least 50% of the cells reacting (deformation and/or agglutination ) was recorded as the endpoint 9 Test 2 Laboratory Transmission
To investigate the m vivo interactions of Spiroplasma and Entomoplasma,
Tabanus longiuscuhs and T. Hneola were captured live, sorted by species, and placed
in sterile 250 mL glass jars containing a 42.5 mm #1 round Whatman® filter paper in
the bottom A plastic petri dish with a 2 mm hole in the center for feeding was used as
a lid for each jar.
The flies were offered a daily drop of 5% sucrose and maintained at an ambient
temperature range of 22-250C The flies were assigned at random to a treatment
group: (A) spiroplasmas only, (B) entomoplasmas only, (C) spiroplasmas first then
entomoplasmas, (D) entomoplasmas first then spiroplasmas, (E) 5% sucrose (control)
Viable Spiroplasma lilorale and Entomoplasma ellychniae cultures in 5%
sucrose were offered in 30 pL drops to the appropriate treatment groups The S
htorale culture (001475) was initially recovered from the abdominal viscera of T. americanus captured in Bulloch County, Georgia during February, 1996 DF testing
resulted in a reaction to TN-1 antisera at a dilution of 1:1280 The E. ellychniae culture (003932) was initially recovered from the abdominal viscera of Hyhomitra tlifficilis captured in Bulloch County, Georgia during April, 1993 DF testing resulted in a reaction to ELCN-1 antisera at a dilution of 1 80
To prepare inoculum, S. litorale and E ellychniae subcultures were removed from a -70oC freezer, rapidly thawed in a warm water bath, 1.5 mL R2 medium added, subcultured, and incubated at 30oC Once a subculture reached the logarithmic growth 10 phase, it was viewed with darkfield microscopy at lOOOx to examine morphology
Tenfold dilution series were prepared in R2 medium ending with dilution 1:1280 The dilutions were viewed with darkfield microscopy at 1500x to estimate the concentration and select the dosage for treatment groups The selected dilution was further diluted by 1/2 with 10% sucrose for feeding the flies and titrated to estimate the number of organisms administered at feeding
The flies were offered a 30 pL hanging drop of 10^ spiroplasmas per cc in 5% sucrose (treatments A and C), 10^ entomoplasmas per cc in 5% sucrose (treatments B and D), or 5% sucrose (treatment E). On post treatment day-4, treatment C was super-infected by offering a 30 pL hanging drop of 10^ entomoplasmas per cc in 5% sucrose and treatment D was super-infected by offering a 30 pL hanging drop of 105 spiroplasmas per cc in 5% sucrose The flies were offered a 30 pL hanging drop of 5% sucrose daily
Five days after super-infection (day 9 after initial treatment), gut and hemolymph isolations were attempted from each fly (Appendix 1) Deformation and endpoint tests were performed on cultures containing spiroplasmas to identify the species.
Cultures appearing to contain entomoplasmas were serocultured to remove spiroplasmas In a 96 well microtiter plate, 20 pL of each culture was mixed with TN-
1 antisera at dilution 1:10 and allowed to react for one hour The reaction mixture was transferred into 1.5 mL fresh R2 medium and syringe filtered through 0.45 gm to remove deformed spiroplasma cells and agglutination masses Once a seroculture reached the logarithmic growth phase, a metabolism inhibition (Ml) test was performed with ELCN-1 antisera (Taylor-Robinson, 1983, Williamson, 1983, Appendix 3). Results
Test 1 Natural Carriage
Chrysops vittatus Infection Rate
Of the 90 C vittatus processed, 23 (25 6%) carried spiroplasmas Chrysops
vittatus was most frequently a host for group VIII spiroplasmas (86.9%): strain B1357
(7/23 or 30.4%); Sptroplasma syrphidicola, strain EA-1 (5/23 or 21.7%); S.
chrysopicola, strain DF-1 (4/23 or 17 4%), strain TAAS-1 (4/23 or 17 4%) Group
XVIII S. litorale, strain TN-1, was detected in 2 of 23 flies (8 7%) and Group XXXV
strain B2649 was detected in 1 of 23 flies (4.4%)
Tabanus atratus Infection Rate
Of the 47 T. atratus processed, 22 (46 8%) carried spiroplasmas. Tabanus atratus was most frequently a host for group XXXIII S. tabanuJicola represented by
strain TAUS-1 (12 of 22-54.6%) followed by group XXIII S. g/aJiatons, strain TG-1.
(3 of 22-13.6%). The remaining isolations were each represented once (1 of 22-4 6% each): group IV strain PPS-1, group XIV S. corruscae, strain EC-1; group XVIII S.
Ulorale, strain TN-1, group XXXII S helicoides, strain TABS-2; group XXXIV strain
B1901, group XXXV strain B2649 No entomoplasmas were detected from the two tabanid species.
12 13 Test 2 Laboratory Transmission
In treatment A (spiroplasmas only), all 12 of the gut cultures attempted reacted
toTN-1 antisera (dilutions 1:640 to 1:1280) in DF testing One of 12 hemolymph
isolation attempts was successfully cultivated and DF testing resulted in reaction to
TN-1 antisera at a dilution of 1:1280. Entomoplasmas were not isolated by
serocultures of gut or hemolymph samples.
From treatment B (entomoplasmas only), three positive gut isolations of
spiroplasmas were obtained DF testing resulted in reaction to TALS-2 antisera at a
dilution of 1:1280 for each isolation No successful hemolymph cultures or
serocultures were obtained
From treatment C (spiroplasmas first then entomoplasmas), DF testing resulted
in a reaction to TN-1 antisera at a dilution range of 1 320 - 1 2560 for all of the nine
gut cultures attempted One hemolymph sample was successfully cultivated and DF
testing resulted in a reaction to TN-1 antisera at a dilution of 1 2560. Seroculture of
entomoplasmas were obtained from the one hemolymph sample and from five gut
samples. MI testing resulted in final liters of 20480 for each gut culture and 10240 for the hemolymph culture.
From treatment D (entomoplasmas first then spiroplasmas), DF testing resulted in a reaction to TN-1 antisera at a dilution range of 1 640 -1:1280 for all of the seven gut cultures attempted. One hemolymph sample was successfully cultured and DF testing resulted in a reaction to TN-1 antisera at a dilution of 1:640, Entomoplasmas 14 were isolated by seroculture from seven gut samples and from the hemolymph sample
MI testing resulted in final titers ranging from 5120 - 20480 for the gut samples and
10240 for the hemolymph sample
From treatment E (control), no S. litorale or A. ellychniae cultures were obtained from the nine gut and nine hemolymph isolations attempted Two samples contained spiroplasmas with a DF test reaction to TALS-2 antisera at dilutions 1 320 and 1 1280 There were no successful serocultures of entomoplasma. Discussion
Test 1 Natural Carriage
Chrysops vittatus, a deer fly, and T atratus, a horse fly, are two of the many tabanid flies that become infected with spiroplasmas. Although they both fly during the summer months and may be captured concurrently in traps, they are infected with different assemblages of spiroplasma species with little overlap In this study, the only common species of spiroplasma to both flies were Spiroplasma htorale and strain
B2649, yet their prevalences were less than 10% of the carriage by each fly species
(Table 1). These results suggest that the two tabanid species acquire spiroplasmas differently.
A similar study was conducted by Moulder, 1997, with female C. brimleyi and
Hybomhra difficihs with an active flight period of April through early May in Bulloch
County, Georgia. The results were similar in that carriage of common species of spiroplasma was also rare (Table 2)
Tabanus lineola is a horse fly whose active flight period in Bulloch County,
Georgia coincides with that of C. vittatus and T atratus French et a/., 1996, conducted a study where the spiroplasma flora diversity of T. lineola was analyzed
Comparison of the results of my investigation with those of French et al., 1996 and
Moulder, 1997 indicates that the two Tabanus spp. carried similar assemblages of spiroplasmas as well as the two Chrysops spp. carried similar assemblages of 15 16 spiroplasmas. The two Chrysops spp carriages were not similar to that of either
Tabanns spp. or H. difficilis (Table 2).
The diversity of spiroplasma assemblages among the tabanids suggests different methods of spiroplasma acquisition. Since there is little overlap in common spiroplasma species, ecological host specificity is also implied as host specificity is not indicated in the laboratory. By investigating the spiroplasma assemblages that tabanids carry, we further our understanding of tabanid ecology as it relates specifically to the acquisition and transmission of pathogens.
Test 2 Laboratory Transmission
Tabanids show no indication in the laboratory of host specificity as they transmit Spiroplasma montanense and S corruscae to one another and readily acquire spiroplasmas by feeding on inoculated sugar water (Railey and French, 1990; French et al., 1994; Wedincamp et a/., 1997). My results show that, for the first time, S litorale is also readily transmitted to tabanids in the laboratory After feeding on sugar water containing S. litorale, all 28 Tabanits lineola and T longiuscuhs acquired the spiroplasmas in treatments A, C, and D (Table 3, Figure 1), Spiroplasma litorale was considered uncommon and restricted to the coastal areas of the southeastern United
States by Konai et al., 1997 as natural infections of A', litorale represented less than 5% of more than 300 isolations from tabanid guts in Bulloch County, Georgia (French et al., 1992, 1996; Konaie/a/., 1997, Moulder, 1997; Table 2). 17 To date, natural infections of/f ellychniae have not been detected or reported from tabanid guts and hemolymph This may be the result of the difficulties arising during microscopic examinations. Due to their pleomorphic capabilities and small size, entomoplasmas are either overlooked or mistaken for other bacteria Although natural infections have not been found, transmission of E. ellychniae from fireflies to tabanids has occurred during cohabitation (Wedincamp et ai, 1996)
Entomoplasma ellychniae was recovered from 5/9 of the gut cultures attempted in treatment C (spiroplasmas first then entomoplasmas) and from 7/7 of the gut cultures attempted in treatment D (entomoplasmas first then spiroplasmas) Overall, E ellychniae was recovered from 50% of 24 gut samples from flies of treatments B, C, and D (all of which were exposed to entomoplasmas) (Table 3; Table 4).
Although treatments B (entomoplasmas only) and D (entomoplasmas first then spiroplasmas) were concurrently offered the same culture and dosage of E. ellychniae. there was no recovery of entomoplasmas from treatment B These results were unexpected because other treatment groups (C and D) were infected using identical methodology. In fact, treatment B and D were offered the same inoculum within five minutes of each other. The most plausible reason is that the flies did not feed, possibly due to a slight difference in overhead lighting
Previous investigators have had minimal success in cultivating mollicutes from tabanid hemolymph (Clark et ai, 1984). My hypothesis was that after a spiroplasma or entomoplasma was allowed to establish a population in the gut, the introduction of a 18 second, different mollicute would promote invasion of the hemolymph Dual infections by S. htorale and E. ellychniae in the hemolymph occurred in 2 of 16 flies (one each in treatments C and D) This is the first report of concurrent infections of S htorale and
E ellychniae in the hemolymph Spiroplasma htorale was recovered from the hemolymph of 1 of 12 tabanids in treatment A with S htorale established in the gut without E. ellychniae. This is the first report of A' htorale isolated from hemolymph.
The frequency of hemolymph infection was less than 10% than that of gut infections
(Figure 1).
The cultures from treatment E (control) were presumed natural infections of A lineolae (strain TALS-2). Overall, in the laboratory, 1 was able to establish the following mollicute populations in tabanids for up to eight (8) days: S. litorate in the gut of 28 tabanids and in the hemolymph of three (3) tabanids, E ellychniae in the gut of 12 tabanids and in the hemolymph of two (2) tabanids; and concurrent infections of
S. htorale and E. ellychniae in the gut of 12 tabanids and in the hemolymph of two (2) tabanids. Literature Cited
Albalain-Colloc, M L., C Chastel, J.G Tully, J M Bove, R.F. Whitcomb, B. Gilot, and
D L Williamson 1987. Spiroplasma sabaudieme, sp nov., a new species from mosquitoes collected in France Int. J Syst Bacteriol 37:260-265
Albalain-Colloc, M L., L Rosen, J G Tully, J M Bove, C Chastel, and D L
Williamson 1988 Spiroplasma taiwanense, sp. nov from Culex tntaenoirhynchus mosquitoes collected in Taiwan Int. J Syst Bacteriol 38:103-107
Bove, J M 1997 Spiroplasmas: Infectious agents of plants, arthropods and vertebrates Wien Klin Wochenschr 14-15:604-612
Chastel, C., B Gilot, F. Le Goff, R. Gruffaz, and M L Albalain-Colloc 1985
Isolement de spiroplasmes en France (Savoire, Alpes du Nord) a partir de Moustiques de genre Aedes Compt Rend Acad Sci (Paris) 300:261-266
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Bacteriol. 40:160-164 Table 1. Spiroplasmas isolated from the abdominal viscera of Chrysops vittatus and
Tabanus atratus captured in Bulloch County, Georgia
Spiroplasma Spiroplasma Hosts group strain species Chrysops vtltalus Tabanus atratus % of n=23 % of n=22 IV PPS-1 not named 0 4 6 VIII DF-1 chrysopicola 17 4 0 VIII EA-1 syrphidicola 21.7 0 VIII TAAS-1 not named 17 4 0 VIII B1357 not named 30.4 0 XIV EC-l cor ru scat' 0 4.6 XVIII TN-1 lilorale 8 7 4 6 XXIII TG-1 gladiatoris 0 13 6 XXXII TABS-2 helicoides 0 4.6 XXXIII TAUS-1 tabanidico/a 0 54.6 XXXIV B1901 not named 0 4 6 XXXV B2649 not named 4.4 4 6
29 30
>5 "d* m II s c a C4- o o o o o o o o o O O § O rj ■P3 vO §■ ov
-c O ri Cs II r Si c 00 ir.0 ^ ' ' 1" T <4- 0 ^r^ 00 — o C3 o -o nO cd "O '5b '•< n (U u rj Urn O o "♦X"- II a »«» 5 120 r 100 80 + 60 40 20 0 t A B C D Treatment Groups Figure 1. Successful isolation attempts from each treatment group. 32 Appendix 1 Isolation Procedures" Gut Isolation To process the gut, each fly was surface sterilized in 20 mL of 0.5% NaOCI containing 2 drops of Photo-Flo®, a surfactant which allows the surface of the fly to become completely wet. After 1 minute, the fly was rinsed in 20 mL of distilled water The fly was then placed on a dry, unbleached paper towel using sterile forceps and continually repositioned until the exterior of the fly was dry The last two segments of the abdomen were removed with a heat sterilized scalpel The visceral contents were then removed with heat sterilized microdissection forceps and placed in a sterile disposable petri dish containing 1 5 mL of M1D or R2 broth. The viscera were minced with a teasing needle and forceps Using a 3 cc syringe and a 16 gauge 1 1/2 " needle, the tissue fragments and broth were removed from the petri dish The mixture was aspirated into the syringe and expelled 3 times to adequately mince the viscera 33 34 The culture was syringe filtered into a sterile 3 mL snap cap culture tube through a 0.45 pm filter and incubated at 30oC for growth Hemolvmph Isolation Before hemolymph samples could be attained, micropipettes were made by heating the tip of a 5 inch Pasteur pipette with a microburner and then drawing it out to approximately 8 inches Each fly was anesthesized using C02 gas With the aid of a dissecting microscope, one fore leg was then removed from the fly with sterile forceps The tip of the micropipette was slightly inserted into the cavity and hemolymph was extracted by capillary action The tip of the micropipette was placed in a 3 mL snap cap culture tube containing 1.5 mL of MID or R2 broth The hemolymph was transferred by gently blowing into the micropipette. The culture was incubated at 30oC for growth Culture Growth and Storage Both gut and hemolymph cultures were monitored daily for growth which is indicated by a gradual change in the color of the broth medium from red to yellow Growth is then confirmed by using darkfield microscopy at lOOOx. 35 If any other fungal or bacterial growth was present, the culture was filtered through a 0.22 pm filter unit attached to a 3 cc syringe The culture is incubated again for growth All confirmed, positive cultures were subcultured and preserved by freezing at - 70oC All insect hosts were labeled with a 6 digit accession number, identified to species, and curated as voucher specimens in the Georgia Southern University insect collection. a (adapted from Wedincamp et al , 1996) Appendix 2 Preparation of R2 Medium*' Combine the following and autoclave at 120oC for 20 minutes: Distilled water 760 mL PPLO broth : 15.0 g Sucrose : 80,0 g Phenol Red (0 2%) : 10 0 mL Allow this mixture to cool to room temperature Transfer 150 mL Horse serum with a sterile, disposable pipette from original bottle into sterile container with a top or lid Heat treat in a 50oC water bath for 30 minutes, stirring frequently Add to cooled, autoclaved mixture Add 1 g / 1000 mL Penicillin G (1 million units/gram) to mixture Mix with magnetic stirrer until completely blended Vacuum filter medium through a 0 22 or 0 45 pin membrane into sterile containers Maintain medium in a contamination-free environment a (adapted from C.J Chang, unpublished data; Moulder, 1997) 36 Appendix 3 Serological Testing Deformation and Endpoint Tests for Spiroolasmas a Cultures to be tested were removed from the -70oC freezer and rapidly thawed in warm water New subcultures were made and incubated at 30oC When the cultures reached the logarithmic phase of growth, they were tested against 6 different combined antiserum (Appendix 4) In a microtiter plate, equal aliquots of combined antisera and culture were mixed and allowed to react for a minimum of 30 minutes The mixture was viewed under darkfield microscopy at lOOOx A positive reaction was indicated by morphological deformities in the helical shape of the spiroplasma or by agglutination of several spiroplasmas For each combined antisera with which the culture reacted, an endpoint test was performed to identify the species of spiroplasma Serial dilutions of the individual antisera components of the combined antisera ranging from 1:10 to 1 2560 were prepared. In a microtiter plate, equal aliquots of culture and serial dilution were mixed and allowed to react for a minimum of 30 minutes. The mixture was viewed under darkfield microscopy at lOOOx The last dilution to denote a 50% reaction (deformation or agglutination) was recorded as the endpoint 37 38 Metabolism Inhibition Test ^ For antigen titration Stock cultures should be titrated to determine the number of viable organisms A total of 12 ten-fold dilutions of the culture are made in complete medium. In a 96 well microtiter plate, 50 pL of each dilution is added to fill one row per culture Six drops (25 pL/drop) of medium is added to each well to produce a total volume of 200 pL The plate is sealed by incubating at 370C for 20 - 30 minutes, then rapidly sealing with 5 cm wide clear commercial package sealing tape at room temperature This prevents loosening of the tape when plates are finally incubated The highest dilution to change color (i.e., if well 10"' through 10"6 change but not 10-7) js regarded as 10^ CCU (CCU means color changing unit) Preparation of 96 - well microtiter plate 1. Place 100 pL of growth medium in wells 1-9, 11, and 12 (antigen titration wells), and 150 pL in well 10 (medium control well) 2. Add 50 pL of 1:10 antiserum dilution in well 1, rows A-H, producing a 1 80 initial antiserum dilution in well 1 3 Serially dilute the antiserum by mixing the antiserum and medium in the first well and transfer 50 pL serially through well 9, 4. Add 50 pL of the antigen to wells 1-9, 11, and 12 5. Add 50 pL of 8% guinea pig complement solution in growth medium to each well 39 6 Cover plate with 5 cm wide clear commercial package sealing tape and incubated at 30oC Ml Titer Determination 1 Monitor the microtiter plate daily for a pH shift indicated by a color change from red to yellow (a decrease in 1 - 1 5 pH units) The highest dilution of antigen giving a color change is considered as one CCU per 50 pL 2 The MI titer is the reciprocal of the highest three-fold antiserum dilution that prevented color change a (adapted from Williamson and Whitcomb, 1978) b (adapted from Williamson, 1983) Appendix 4 Six Different Combined Antisera for Screening Combination Group Scientific Name Strain IV Spiroplastna apis B31 IV not named W13 IV not named PPS-1 VIII-1 S. syrphic/ico/a EA-1 VIII-2 S. chrysopicola DF-1 VIII-3 not named TAAS-i VIII not named B1357 XIV S. corniscae EC-1 XXXI S. montanense HYOS XXIII S. gladialoris TG-1 XXXIII S. tabanidicoki TAUS- XXVIII S. hneola TALS-2 XVIII S. htorale TN-1 XXXII S. helicoides TABS-2 XXXIV not named B1901 XXXV not named B2649 40