Attempted Mechanical Transfer of Ehrlichia risticii by Tabanids (Diptera: Tabanidae) JAY F. L E V I N E , M I C H A E L G. LEVY, WILLIAM L. NICHOLSON, WILLIAM S. IRBY, ROBIN GAGER, AND CHARLES S. APPERSON 1 Department of Microbiology, Pathology, and Parasitology, College of Veterinary Medicine, North Carolina State University, Raleigh, North Carolina 27606
KEY WORDS Insecta, Tabanidae, Ehrlichia risticii, mechanical transfer
POTOMAC HORSE FEVER, equine monocytic ehrlichiosis (EME), was initially diagnosed in the vicinity of the Potomac River in the late 1970s (Knowles et al. 1984). The etiologic agent of EME, Ehrlichia risticii Holland, Weiss, Burgdorfer, Cole & Kakoma, was recognized as a new species in the genus Ehrlichia (Holland et al. 1985a, Rikihisa & Perry 1985). Experimental studies have confirmed the association of the disease with this agent, which rapidly produces clinical disease in susceptible horses (Dutta et al. 1985, Holland et al. 1985b, Perry et al. 1985). Equine monocytic ehrlichiosis is generally diagnosed from June through September (Gordon et al. 1988); this seasonal distribution suggests that arthropods play a role in the maintenance of E. risticii transmission (Carroll & Schmidtmann 1986, Schmidtmann et al. 1986). Various arthropod pests of horses are present on farms where EME has been diagnosed, and several species have been suggested as vectors. Initial studies focused on the ixodid tick Dermacentor variabilis (Say); however, attempts to transmit the pathogen with D. variabilis (Schmidtmann et al. 1986, Hahn et al. 1990, Levine et al. 1990) and other ixodid ticks (Hahn et al. 1990) were unsuccessful. Several genera of biting flies, including Tabanus, Chrysops, and Stomoxys, have been temporally and geographically associated with 1 Department of Entomology, College of Agriculture and Life Sciences, North Carolina State University, Raleigh, NC 27695.
EME (Fletcher et al. 1988, Burg et al. 1990, Burg et al. 1991). Recognition of the principal vector of E. risticii is a prerequisite to development of effective strategies to control spread of the disease. Accordingly, these studies were initiated to evaluate the potential of various species of tabanid biting flies to serve as mechanical vectors of E. risticii.
Materials and Methods Sampling Sites and Fly Sampling Methods. Horse flies and deer flies have not been colonized in the laboratory. Consequently, tabanids used in this study were field-collected specimens. Flies were collected primarily at four sites in Wake and Johnston counties of North Carolina: (1) a commercial boarding stable and riding school in northwest Wake County, NC; (2) William B. Umstead State Park, a 2,024-ha wildlife and recreational facility adjacent to Raleigh, NC, in Wake County; (3) a commercial boarding stable and riding academy southwest of Raleigh, NC, in Wake County; and (4) a private residence near Clayton, NC, in Johnston County. Limited sampling was conducted at two additional sites in Wake County and at one site in Duplin County to supply additional specimens for testing. All sites had a history of biting fly activity and resident or transient horse activity. No cases of EME had been reported at these sites.
0022-2585/92/0806-0812$02.00/0 © 1992 Entomological Society of America
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
J. Med. Entomol. 29(5): 806-812 (1992) ABSTRACT The ability of tabanid mouthparts to retain and to transfer mechanically Ehrlichia risticii Holland, Weiss, Burgdorfer, Cole & Kakoma was evaluated by feeding flies on infected and noninfected mice and on capillary tubes containing infected cells and cell-free medium. Flies representing two genera and 29 species were collected at equine boarding stables, farms, and along riding trails in Wake, Johnston, and Duplin counties in North Carolina for the feeding trials. Two species, Tabanus fulvulus Wiedemann and T. pallidescens Philip, fed on mice but failed to transfer the pathogen from infected to susceptible mice. Chrysops vittatus Wiedemann, Tabanus americanus Forster, and T. sulcifrons Macquart transferred E. mticu-infected cells from capillary tubes containing infected cells in medium to tubes containing medium. These studies document that £. nsticu-infected cells can be retained on mouthparts and potentially transferred by tabanids.
September 1992
LEVINE ET AL.: MECHANICAL TRANSFER OF E. risticii BY TABANIDS
suspended in medium and the cell number determined with a cell-counting chamber. Feeding on a Murine Host. Flies were individually fed on the shaven back of a Balb/C mouse anesthetized with ketamine hydrochloride (50 mg/kg) and xylazine (20 mg/kg). Mice were inoculated with 105 E. risticii-iniected cells intraperitoneally each week, and showed clinical signs of infection when used as hosts 7—10 d after inoculation. For each trial, a fly was confined to the back of each mouse by a small ventilated polystyrene cup taped to the mouse and watched for 10—15 min for evidence of probing or other signs of feeding. If blood ingestion occurred, feeding was interrupted after 2—3 min, and theflywas transferred to the shaven back of an uninfected, fly-naive mouse. Flies were then allowed to feed undisturbed until they were engorged or the trial was terminated. Flies that did not feed initially on mice were retested on mice after being deprived of sucrose. Flies were also retested using the capillary tube method. Mice that were fed upon or probed by the flies were monitored for evidence of exposure to E. risticii. Mice were bled weekly for 6 wk from the tail and the plasma frozen at — 20°C until it was assayed. The plasma was tested for antibody to E. risticii by an indirect immunofluorescence assay (IFA) (Rikihisa et al. 1987). Plasma was initially tested at a 1/20 dilution. Positive and negative controls were included on each slide. Antigen-containing slides were prepared by growing E. risticii in P388-DJ cells, as described above, until the cells were >80% infected. The cells were dislodged with a cell scraper and resuspended by shaking. The suspension was centrifuged at 1,500 x gfor10 min at 4°C. The pellet was washed once in RPMI 1640 medium and resuspended in 0.1 M phosphate-buffered saline (pH 7.3) containing 0.5% (wt/vol) bovine serum albumin (PBS-BSA). The suspension was dotted onto 24-well template slides (Cell-Line Associates, Newfield, NJ), air dried, and fixed in acetone for 10 min. Prepared slides were stored at -20°C until they were tested by IFA. Before testing, antigen-containing slides were washed once with water, the wells covered with 10% (vol/vol) normal goat serum in PBS, placed in a humidified chamber, and incubated for 30 min at 37°C. After a 5-min wash in PBS, the wells were covered with dilutions of the mouse plasma to be tested made in PBS-BSA, and incubated for 30 min at 37°C. Slides were then washed four times (10 min each) in PBS. A 1/50 dilution of FITC-labeled goat-antimouse IgG (H+L) (Cat. no. 1211-0081, Organon Teknika Corp., Durham, NC) in PBS-BSA was applied to each well, incubated for 30 min at 37°C, and washed in PBS four times (10 min each). In the last wash, 0.1-0.2 ml 0.125% (wt/vol) Evans Blue per 20-40 ml of wash was added to counterstain the P388-D! cells. Slides were dried and coverslips affixed to
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
Fly trapping was conducted from May through the first week in October 1988. Female tabanids were collected with canopy traps with black ball decoys (Catts 1970), supplemented with carbon dioxide (dry ice) as an attractant (Everett & Lancaster 1968). Flies were funneled into a screened collection chamber at the apex of the trap. Aerial netting was used to supplement canopy trap collections (Tallamy et al. 1976). Additional specimens were collected as they entered our vehicle. Flies were transported to the laboratory in insulated containers cooled with frozen gel packs and were held at room temperature until used in the feeding trials. All flies collected were identified with dichotomous keys (Pechuman 1973, Drees et al. 1980); identifications were verified by L. L. Pechuman, Cornell University, Ithaca, NY. Voucher specimens were deposited in the Insect Collection of the Department of Entomology, North Carolina State University. Mice. Adult female Balb/C (Charles Rivers Laboratories, Raleigh, NC) mice were acclimated to the laboratory for at least 1 wk before the experiments were begun. The mice were kept in plastic cages in an Illinois-type ventilated chamber at ambient temperature (~25°C). Mice were provided with water and standard rodent chow throughout the study. Inoculum Preparation. Ehrlichia risticii (obtained from Y. Rikihisa of Ohio State University and C. Pretzman, Ohio Department of Health) was propagated in the continuous murine macrophage cell line P388-Di (ATCC TIB 63, American Type Culture Collection, Rockville, MD) as previously described (Pretzman et al. 1987). We previously found the strain to be pathogenic to both mice and horses, with no loss in infectivity due to passage (unpublished data). The P388-D! cell line was propagated in RPMI 1640 medium (Gibco Laboratories, Grand Island, NY) supplemented with 10% fetal bovine serum held at 37°C in an atmosphere of 5% carbon dioxide in air. When a confluent cell layer was attained, the medium was removed and ^ l O 3 E. risticiiinfected P388-D! cells in 1-2 ml of medium were added to the flask. The flasks were incubated for several hours at 35°C with periodic rocking to distribute the infected cell suspension evenly. Additional fresh medium (4.5 ml) was added to each flask, and the cultures were held at 35°C. Infected cells were harvested for preparation of the inoculum when 95% of the cells became infected with E. risticii, as determined by cytocentrifugation of the nonadherent cells and staining with a triarylmethane—xanthene—thiazine stain (Diff-Quik, American Scientific Products, McGraw Park, IL). The confluent cells were disturbed with a cell scraper and resuspended by shaking. The cells were centrifuged at 1,500 x g for 10 min at 4°C, and the pellet was washed once in RPMI medium. The final pellet was re-
807
808
Vol. 29, no. 5
JOURNAL OF MEDICAL ENTOMOLOGY
the slides with PBS-glycerol mounting medium. The preparation was examined at 250 x under epifluorescence microscopy for specific ehrlichial fluorescence. Feeding on Medium in Capillary Tubes.
Genus Tabanus
Genus Chrypsops
T. americanus Forster T. atratus atratus F. T. fulvulus Wiedemann T. gladiator Stone T. imitans Walker T. lineola F. T. melanocerus Weidemann T. molestus molestus Say T. nigrescens Palisot de Beauvois T. nigripes Wiedemann T. pallidescens Philip T. petiolatus Hine T. quinquevittatus Wiedemann T. sparus milleri Whitney T. subsimilis Bellardi T. sulcifrons Macquart T. trimaculatus Palisot de Beauvois
C. callidus Osten Sacken C. celatus Pechuman C. dacne Philip C. flavidus Wiedemann C. geminatus Wiedemann C. macquarti Philip C. montanus Osten Sacken C. pikei Whitney C. reicherti Fairchild C. univittatus Macquart C. upsilon Philip C. vittatus Wiedemann
medium. The second was then removed and tapped onto teflon template-printed slides, air dried, and fixed in acetone for 10 min. Slides were examined as previously described. Results
Tabanids representing two genera and 29 species were collected (Table 1). Although 83 tabanids were tested by feeding on E. risticiiinfected mice (Table 2), only T. fulvulus and T. pallidescens fed upon the mice. Of the 13 flies that probed infected mice, seven ingested a partial or full blood meal. All 13 flies were transferred to uninfected mice, and 7 took partial or full blood meals. Three of the flies that obtained a partial meal on the first host did not attempt to feed on the second host. Three flies that probed but did not ingest blood from the first host fed to repletion on the second mouse. During the 6-wk monitoring period following fly feeding, no antibody to E. risticii was detected in the plasma of the second group of mice, which were probed by tabanids that had previously probed on infected mice. Flies readily fed on medium in capillary tubes, ingesting from 0.5 to nearly 30 fi\ of medium per fly (Table 3). The amount ingested varied with the size of the fly, but even small flies were capable of taking in relatively large volumes of fluid. Intermittent contractions of the abdomen were evident throughout the feeding trials, and, occasionally, a droplet of clearfluidwas expelled from the anus of the fly. Standard capillary tubes worked well with a wide range of flies; for those with large mouthparts, larger tubes were necessary. The ability of 8 species of Chrysops and 12 species of Tabanus to retain E. risticii on their mouthparts was tested by the capillary tube
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
Horse flies and deer flies were offered infectious meals in a manner similar to that previously described for ixodid ticks (Burgdorfer 1957). Each fly was placed on its dorsal surface and held down by strands of clay. An "arm" of clay supported the capillary tube around the mouthparts of the flies. Standard plain (nonheparinized) glass capillary tubes (1.1-1.2 mm inside diameter) or larger laboratory-prepared tubes (1.5 to 1.8 mm inside diameter) were dipped into a suspension of 1056 10 /ml E. risticii-inkcted P388-Di cells in RPMI 1640 medium containing 10% FBS. The tube was placed over the mouthparts of each fly and supported by the clay arm. Flies were allowed to ingest the suspension for 5 min, the tube was removed, and the flies were held without feeding for 2-5 min. A tube of RPMI 1640 with 10% FBS (no cells) was then lowered over the mouthparts, and the flies were allowed to ingest fluid from the second tube for another 5 min. The volumes offluidingested were recorded for each tube. The contents of the second feeding tubes were initially examined by cytocentrifugation and Diff-Quik (American Scientific Products, McGraw Park, IL) staining. This technique proved unsuitable, and any transferred cells were subsequently detected by an IFA using hyperimmune mouse anti-E. risticii serum. The contents of both tubes were dotted onto the wells of a 30-well printed glass slide, air-dried, and acetone-fixed to the slide. The assay was conducted as previously described above, except that the test serum was replaced with a 1/100 dilution of hyperimmune mouse serum, which served as the antigen detection probe. Hyperimmune serum (IFA titer of 1/2,560) was collected from mice experimentally infected with E. risticii. The entire area of each well (2 mm diameter) was searched at 250 x with epifluorescence microscopy for specific fluorescence of organisms within the cell cytoplasm. Fluorescent particles not associated with cells were not considered ehrlichial organisms for this assay. Positive and negative controls were included in each batch of slides tested. The ability to transfer cells on the surface of a needle was also examined using the same protocol used to evaluate transfer of E. risticii between capillary tubes by flies. Hypodermic needles (22 gauge) were cut to leave a 5-mm shaft. Cell suspensions containing E. risticii in capillary tubes were applied to the needles. The capillary tubes containing marked cells were replaced with a capillary tube containing cell-free
Table 1. Tabanids collected at seven sites in Wake (n = 5), Johnston (n = 1), and Duplin (n = 1) counties, N.C., during early May—October 1988
September 1992
LEVINE ET AL.: MECHANICAL TRANSFER OF
E. risticii BY
809
TABANIDS
Table 2. Exposure time, probing attempts, and engorgement success of T. fulvulus (Tf) and T. pallidescens placed on £ . rwtieii-infected mice and transferred to uninfected mice. Infected mouse Fly no"
Time on host, min
Probe
Tf-NN Tp-5 Tf-19 Tf-20 Tf-21 Tf-22 Tf-24 Tf-102 Tf-116 Tf-175 Tf-208 Tf-209 Tf-213
2 14 15 7 13 10 8 18 4 18 11 5 9
Y Y Y Y Y Y Y Y Y Y Y Y Y
6
Engorgement levelc
period, min
Time on host, min
P P O O P
0 6 1 4 0 3 2 4 60 0.3 1 0.3 0.2
8 22 13 18 10-15 10-15 10-15 6 8 10 3 15 9
o op o oF P P
(Tp)
Uninfected mouse Engorgement Probe6 levelc N Y Y Y N N N Y Y N N Y Y
O F F F O O O P F O O P P
method (Table 3). Ten of 109 (9.2%) flies retained cells on their mouthparts after feeding on medium containing infected cells and transferred the cells to medium in a second tube. No E. risticii were observed in the cells transferred by two flies. Cell transfer was observed in 10 of 10 trials when needles were used as substrate. Discussion Demonstration of the competence of arthropods as vectors requires recognition of infection
in a host after transfer of the agent by the arthropod. Flies that probed on £. mticii-infected mice apparently failed to transfer an infectious dose of the agent to noninfected mice. Previous to this study, we confirmed the validity of the IFA procedure for detection of antibody to E. risticii in murine serum (unpublished data), and IFA provided a viable means of assessing the exposure of mice to E. risticii. Although it is unlikely that mouse-to-mouse transmission by flies occurred, a small number of infected cells may have been transferred but were insufficient
Table 3. Mean volume (±SD) of infected cell suspension and medium ingested in 5 min by tabanids individually fed by the capillary tube method, and transfer of cells as determined by immunofluorescence assay Species Chrysops spp. C. callidus C. celatus C. dacne C.flavidus C. geminatus C. montanus C. univittatus C. vittatus
C. spp.
Tabanusspp. T. americanus T. atratus atratus T. fuloulus T. lineola T. melanocerus T. nigrescens T. nigripes T. petiolatus T. quinquevittatus T. sparus milleri T. subsimilis T. sulcifrons T. spp. Undetermined tabanids Total
No.
2 5 2 5 1 1 1 30 33 3 1 1 1 3 1 1 1 3 9 3 17 3 7 134
° ND, not done. b Larger laboratory-prepared tubes were used.
Vol cell suspension, fi\ 1.0 3.3 2.0 4.6 1.0 1.0 3.0 2.5 1.7
(2.6) (5.7)
(2.1) (1.5)
9.8 (5.0) 28.0b 3.5 16.0 6.0 (6.2) 1.0 6.0 2.0 2.3 (1.2) 3.4 (2.8) 1.7 (1.2) 7.2 (5.7) 8.2 (6.9) 7.5 (9.2)
Vol medium, /xl 3.0 1.4 0.8 3.3 1.0 0.5 0 3.3 1.6
(1.1) (6.0)
(2.9) (1.3)
3.0 (2.5) 12.4fc 4.0 5.0 1.8 (0.8) 1.0 2.0 1.5 1.0 (0.5) 2.6 (1.5) 0.8 (0.3) 11.0 (13.3) 3.8 (1.3) 2.9 (1.9)
No. positive/ total tested" 0/ 2 0/ 4 ND 0/ 4 ND 0/ 1 0/ 1 1/26 (3.8%) 3/25 (12%) 1/ 3 (33.3%) 0/ 1 ND 0/ 0 0/ 3 0/ 1 0/ 1 0/ 1 0/ 3 0/ 7 0/ 3 4/14 (28.6%) 0/ 3 1/ 6 (16.7%) 10/109 (9.2%)
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
" Tf, Tabanus fulvulus; Tp, Tabanus pallidescens. b Probe success: Y, yes; N, no. c Engorgement level: O, no visible abdomen expansion; P, partial blood meal taken; F, full blood meal taken.
810
JOURNAL OF MEDICAL ENTOMOLOGY
of infected cells that can potentially be ingested or contaminate the mouthparts of a fly during feeding. Actual volumes of blood and tissue fluid ingested, however, will vary with the species of fly feeding. Foil et al. (1989), in studies conducted with T.fusciostatus and bovine leukemia virus, found that as few as 10fliestransferred that virus from infected to noninfected hosts. The authors estimated that 10flieswould move up to 60 /Ltl of infected blood between hosts. The actual potential for transmission is also dependent on the number of infected cells present at the time of feeding (Foil et al. 1989). Precise verification of the ehrlichemia developed by infected horses and the actual number of E. risticii cells retained in bloodmeal residue on fly mouthparts is needed to quantify the potential for tabanid transmission accurately. Mechanical transmission occurs when a fly feeding on a host with a patent infection is interrupted and the arthropod moves to a susceptible host. The vector potential of each fly species for a given pathogen, however, is quite variable and is dependent on: (1) the stability of the agent on fly mouthparts, (2) the quantity of infected material that can be transferred between hosts, and (3) the abundance and biting habits of individual fly species (Foil 1989). Host skin thickness, the response of the host to fly biting, and the maintenance of a patent infection determine the hosts' ability to support mechanical transmission by a competent vector (Foil et al. 1985, Foil 1989). The spatial and temporal distribution of immune and susceptible hosts also affect the potential for mechanical transfer of an agent. Epidemiologic studies at locations where equine monocytic ehrlichiosis has been diagnosed suggest that cases are generally spatially isolated (Palmer et al. 1986, Burg et al. 1990) and that mechanical transmission of E. risticii between horses might be unlikely. However, in one study, antibody to E. risticii was detected in the serum of cats residing on farms where cases of EME had been observed (Perry et al. 1989). Horses may be dead-end hosts (Burg et al. 1990), and the role of cats or other hosts as potential reservoirs that help support E. risticii transmission may warrant additional study (Perry et al. 1989). Our studies demonstrate that E. risticiiinfected cells can be retained on the mouthparts of some species of tabanids. However, additional experiments with infected and susceptible horses and possibly other hosts are needed to document the vector-competence of tabanids and the capacity of individual species to maintain transmission after the pathogen is introduced into an equine population. Acknowledgment We thank L. L. Pechuman (Cornell University) for confirming the identity of the tabanids, D. Moncol and
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
to induce an antibody response. Isolation of E. risticii would have provided a direct alternative measure of infection, but it was not attempted. Three species of tabanids fed by the capillary tube method retained cells on their mouthparts and transferred infected cells between the capillary tubes, demonstrating that E. risticiiinfected cells can be successfully carried on the mouthparts of tabanids. Studies conducted with T.fuscicostatus have shown that ^lO nl of residual blood remain on the mouthparts of this species when feeding on a host is interrupted (Foil et al. 1987). The period of time that an agent remains viable on the mouthparts offliesis quite variable and is dependent on related factors (pathogen, environmental, vector) (Foil 1989). We used an interfeed period of 2—5 min between capillary tubes, and E. m£icu-infected cells remained intact on the mouthparts of at least three species during this period. Bloodmeal residue has been detected on tabanid mouthparts hours after the interruption of feeding, but detectable residues decline significantly after several hours (Foil et al. 1987). Fly grooming and mouthpart cleaning may account for some reduction in residues. Tabanids are recognized mechanical vectors of >30 pathogens that cause disease in animals (Krinsky 1976). Tabanid biting activity can be intense. In some areas, tabanids have been observed landing on horses at a rate of 1,000 flies per hour. Several species of tabanids may serve as mechanical vectors, and transfer of an infective dose may occur through the combined attack of multiple species (Foil 1989). Actual biting rates in a pasture will vary with the species of flies present, the presence of suitable habitat, climate, herd size, animal behavior, and the distribution of animals. Although the ability of numerous species of biting flies to transmit pathogens has been well documented, no Ehrlichia sp. is known to be fly-transmitted. The Ehrlichieae are obligate intracellular parasites (Ristic & Huxsoll 1984), and successful mechanical transmission requires the transfer of intact infected cells between hosts. Tabanids and iatrogenic spread by needle are recognized means of transmission of other cellassociated pathogens such as equine infectious anemia virus (Hawkins et al. 1976, Issel et al. 1988), bovine leukemia virus (Bech-Nielson et al. 1978, Foil et al. 1988), and Anaplasma marginale (Dikmans 1950). These studies suggest that E. risticii cells may remain viable on the mouthparts of some species of tabanids and on the surface of needles. Biologic transmission of E. risticii by Stomoxys calcitrans has also been evaluated. However, this species failed to transfer infection from an infected to a noninfected host (Burg et al. 1990). The concentration of a pathogen in the blood and local tissues of a host determine the number
Vol. 29, no. 5
September 1992
LEVINE ET AL.: MECHANICAL TRANSFER OF E. risticii BY TABANIDS
A. Artis for technical assistance, and E. W. Cupp (University of Arizona) for suggesting the tube-feeding method. Funds to support this research were provided by the Morris Animal Foundation.
References Cited
investigation of farms with Potomac horse fever (equine monocytic ehrlichiosis). Acta Vet. Scand. (Suppl.) 84: 319-322. Hahn, N. E., M. Fletcher, R. M. Rice, K. M. Kocan, J. W. Hansen, J. A. Hair, R. W. Barker & B. D. Perry. 1990. Attempted transmission of Ehrlichia risticii, causative agent of Potomac horse fever, by the ticks, Dermacentor variabilis, Rhipicephalus sanguineus, Ixodes scapularis, and Amblyomma americanum. Exp. Appl. Acarol. 8: 41-50. Hawkins, J. A., W. V. Adams, Jr., B. H. Wilson, C. J. Issel & E. E. Robl. 1976. Transmission of equine infectious anemia virus by Tabanus fuscicostatus. J. Am. Vet. Med. Assoc. 168: 63-64. Holland, C. J., E. Weiss, W. Burgdorfer, A. I. Cole & I. Kakoma. 1985a. Ehrlichia risticii sp. nov.: etiological agent of equine monocytic ehrlichiosis (synonym, Potomac horse fever). Int. J. Syst. Bacteriol. 35: 524-526. Holland, C. J., M. Ristic, A. I. Cole, P. Johnson, G. Baker & T. Goetz. 1985b. Isolation, experimental transmission, and characterization of causative agent of Potomac horse fever. Science 227: 522524. Issel, C. J., K. Rushlow, L. D. Foil & R. C. Montelaro. 1988. A perspective on equine infectious anemia with an emphasis on vector transmission and genetic analysis. Vet. Microbiol. 17: 251-286. Knowles, R. C , D. W. Anderson, W. D. Shipley, R. H. Whitlock, B. D. Perry & J. P. Davidson. 1984. Acute equine diarrhea syndrome (AEDS): a preliminary report. Proc. Am. Assoc. Equine Pract. 29: 353-357. Krinsky, W. L. 1976. Animal disease agents transmitted by horse flies and deer flies (Diptera: Tabanidae). J. Med. Entomol. 13: 225-275. Levine, J. F., M. G. Levy, W. L. Nicholson & R. B. Gager. 1990. Attempted Ehrlichia risticii transmission with Dermacentor variabilis (Acari: Ixodidae). J. Med. Entomol. 27: 931-933. Palmer, J. E., R. H. Whitlock & C. E. Benson. 1986. Equine ehrlichial colitis (Potomac horse fever): recognition of the disease in Pennsylvania, New Jersey, New York, Ohio, Idaho, and Connecticut. J. Am. Vet. Med. Assoc. 189: 197-199. Pechuman, L. L. 1973. Horse flies and deer flies of Virginia (Diptera: Tabanidae). Va. Polytech. Inst. State Univ. Res. Div. Bull. 81. Perry, B. D., Y. Rikihisa & G. K. Saunders. 1985. Intradermal transmission of Potomac horse fever. Vet. Rec. 16: 246-274. Perry, B. D., E. T. Schmidtmann, R. M. Rice, J. W. Hansen, M. Fletcher, E. C. Turner, M. G. Robl & N. E. Hahn. 1989. Epidemiology of Potomac horse fever: an investigation into the possible role of nonequine mammals. Vet. Rec. 125: 83-86. Pretzman, C. I., Y. Rikihisa, D. Ralph, J. C. Gordon & S. Bech-Nielson. 1987. Enzyme-linked immunosorbent assay for Potomac horse fever disease. J. Clin. Microbiol. 25: 31-36. Rikihisa, Y. & B. D. Perry. 1985. Causative ehrlichial organisms in Potomac horse fever. Infect. Immun. 49: 513-517. Rikihisa, Y., G. C. Johnson & C. J. Burger. 1987. Reduced immune responsiveness and lymphoid depletion in mice infected with Ehrlichia risticii. Infect. Immun. 55: 2215-2222. Ristic, M. & D. L. Huxsoll. 1984. Tribe II. Ehr-
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
Bech-Nielsen, S., C. E. Piper & J. F. Ferrer. 1978. Natural mode of transmission of the bovine leukemia virus: role of bloodsucking insects. Am. J. Vet. Res. 39: 1089-1092. Burg, J. G., A. W. Roberts, N. M. Williams, D. G. Powell & F. W. Knapp. 1990. Attempted transmission of Ehrlichia risticii (Rickettsiaceae) with Stomoxys calcitrans (Diptera: Muscidae). J. Med. Entomol. 27: 874-877. Burg, J. G., D. G. Powell & F. W. Knapp. 1991. Arthropod faunal composition on Kentucky equine premises. J. Med. Entomol. 28: 658-662. Burgdorfer, W. 1957. Artificial feeding of ixodid ticks for studies on the transmission of disease agents. J. Infect. Dis. 100: 212-214. Carroll, J. F. & E. T. Schmidtmann. 1986. American dog tick (Acari: Ixodidae) summer activity on equine premises enzootic for Potomac horse fever in south-central Maryland. J. Econ. Entomol. 79: 62-66. Catts, E. P. 1970. A canopy trap for collecting Tabanidae. Mosq. News 30: 472-474. Dikmans, G. 1950. The transmission of anaplasmosis. Am. J. Vet. Res. 11: 5-16. Drees, B. M., L. Butler & L. L. Pechuman. 1980. Horse flies and deer flies of West Virginia: an illustrated key (Diptera, Tabanidae). W. Va. Univ. Agric. For. Exp. Stn. Bull. 674. Dutta, S. K., A. C. Myrup, R. M. Rice, M. G. Robl & R. C. Hammond. 1985. Experimental reproduction of Potomac horse fever in horses with a newly isolated Ehrlichia organism. J. Clin. Microbiol. 22: 265-269. Everett, R. & J. L. Lancaster. 1968. A comparison of animal and dry-ice baited traps for the collection of tabanids. J. Econ. Entomol. 61: 863-864. Fletcher, M. G., E. C. Turner & J. W. Hansen. 1988. Horse-baited insect trap and mobile insect sorting table used in a disease vector identification study. J. Am. Mosq. Control Assoc. 4: 431-435. Foil, L. D. 1989. Tabanids as vectors of disease agents. Parasitol. Today 5: 88-96. Foil, L. D., D. Stage, W. V. Adams, Jr. & C. J. Issel. 1985. Observations of tabanid feeding on mares and foals. Am. J. Vet. Res. 46: 1111-1113. Foil, L. D., W. V. Adams, J. M. McManus & C. J. Issel. 1987. Bloodmeal residue on mouthparts of Tabanus fuscicostatus (Diptera: Tabanidae) and the potential for mechanical transmission of pathogens. J. Med. Entomol. 24: 613-616. Foil, L. D., C. L. Seger, D. D. French, C. J. Issel, J. M. McManus, C. L. Ohrberg & R. T. Ramsey. 1988. Mechanical transmission of bovine leukemia virus by horse flies (Diptera: Tabanidae). J. Med. Entomol. 25: 374-376. Foil, L. D., D. D. French, P. G. Hoyt, C. J. Issel, D. J. Leprince, J. M. McManus & C. L. Seger. 1989. Transmission of bovine leukemia virus by Tabanus fuscicostatus. Am. J. Vet. Res. 50: 1771-1773. Gordon, J. C , S. Bech-Nielson, C. Kohn, W. Farrar, M. Parsons & W. Foster. 1988. An epidemiological
811
812
JOURNAL O F MEDICAL ENTOMOLOGY
lichieae, pp. 704-709. In N. R. Krieg [ed.], Bergey's manual of systematic bacteriology, vol. 1. Williams & Wilkms, Baltimore. Schmidtmann, E. T., M. G. Robl & J. F. Carroll. 1986. Attempted transmission of Ehrlichia risticii by field-captured Dermacentor variabilis (Acari: Ixodidae). Am. J. Vet. Res. 47: 2393-2395.
Vol. 29, no.
5
Tallamy, D. W., E. J. Hansens & R. F. Denno. 1976. A comparison of Malaise trapping and aerial netting for sampling a horse fly and deer fly community. Environ. Entomol. 5: 788-792. Received for publication 2 December 1991; accepted 6 April 1992.
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
KEY WORDS Insecta, Tabanidae, Ehrlichia risticii, mechanical transfer
POTOMAC HORSE FEVER, equine monocytic ehrlichiosis (EME), was initially diagnosed in the vicinity of the Potomac River in the late 1970s (Knowles et al. 1984). The etiologic agent of EME, Ehrlichia risticii Holland, Weiss, Burgdorfer, Cole & Kakoma, was recognized as a new species in the genus Ehrlichia (Holland et al. 1985a, Rikihisa & Perry 1985). Experimental studies have confirmed the association of the disease with this agent, which rapidly produces clinical disease in susceptible horses (Dutta et al. 1985, Holland et al. 1985b, Perry et al. 1985). Equine monocytic ehrlichiosis is generally diagnosed from June through September (Gordon et al. 1988); this seasonal distribution suggests that arthropods play a role in the maintenance of E. risticii transmission (Carroll & Schmidtmann 1986, Schmidtmann et al. 1986). Various arthropod pests of horses are present on farms where EME has been diagnosed, and several species have been suggested as vectors. Initial studies focused on the ixodid tick Dermacentor variabilis (Say); however, attempts to transmit the pathogen with D. variabilis (Schmidtmann et al. 1986, Hahn et al. 1990, Levine et al. 1990) and other ixodid ticks (Hahn et al. 1990) were unsuccessful. Several genera of biting flies, including Tabanus, Chrysops, and Stomoxys, have been temporally and geographically associated with 1 Department of Entomology, College of Agriculture and Life Sciences, North Carolina State University, Raleigh, NC 27695.
EME (Fletcher et al. 1988, Burg et al. 1990, Burg et al. 1991). Recognition of the principal vector of E. risticii is a prerequisite to development of effective strategies to control spread of the disease. Accordingly, these studies were initiated to evaluate the potential of various species of tabanid biting flies to serve as mechanical vectors of E. risticii.
Materials and Methods Sampling Sites and Fly Sampling Methods. Horse flies and deer flies have not been colonized in the laboratory. Consequently, tabanids used in this study were field-collected specimens. Flies were collected primarily at four sites in Wake and Johnston counties of North Carolina: (1) a commercial boarding stable and riding school in northwest Wake County, NC; (2) William B. Umstead State Park, a 2,024-ha wildlife and recreational facility adjacent to Raleigh, NC, in Wake County; (3) a commercial boarding stable and riding academy southwest of Raleigh, NC, in Wake County; and (4) a private residence near Clayton, NC, in Johnston County. Limited sampling was conducted at two additional sites in Wake County and at one site in Duplin County to supply additional specimens for testing. All sites had a history of biting fly activity and resident or transient horse activity. No cases of EME had been reported at these sites.
0022-2585/92/0806-0812$02.00/0 © 1992 Entomological Society of America
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
J. Med. Entomol. 29(5): 806-812 (1992) ABSTRACT The ability of tabanid mouthparts to retain and to transfer mechanically Ehrlichia risticii Holland, Weiss, Burgdorfer, Cole & Kakoma was evaluated by feeding flies on infected and noninfected mice and on capillary tubes containing infected cells and cell-free medium. Flies representing two genera and 29 species were collected at equine boarding stables, farms, and along riding trails in Wake, Johnston, and Duplin counties in North Carolina for the feeding trials. Two species, Tabanus fulvulus Wiedemann and T. pallidescens Philip, fed on mice but failed to transfer the pathogen from infected to susceptible mice. Chrysops vittatus Wiedemann, Tabanus americanus Forster, and T. sulcifrons Macquart transferred E. mticu-infected cells from capillary tubes containing infected cells in medium to tubes containing medium. These studies document that £. nsticu-infected cells can be retained on mouthparts and potentially transferred by tabanids.
September 1992
LEVINE ET AL.: MECHANICAL TRANSFER OF E. risticii BY TABANIDS
suspended in medium and the cell number determined with a cell-counting chamber. Feeding on a Murine Host. Flies were individually fed on the shaven back of a Balb/C mouse anesthetized with ketamine hydrochloride (50 mg/kg) and xylazine (20 mg/kg). Mice were inoculated with 105 E. risticii-iniected cells intraperitoneally each week, and showed clinical signs of infection when used as hosts 7—10 d after inoculation. For each trial, a fly was confined to the back of each mouse by a small ventilated polystyrene cup taped to the mouse and watched for 10—15 min for evidence of probing or other signs of feeding. If blood ingestion occurred, feeding was interrupted after 2—3 min, and theflywas transferred to the shaven back of an uninfected, fly-naive mouse. Flies were then allowed to feed undisturbed until they were engorged or the trial was terminated. Flies that did not feed initially on mice were retested on mice after being deprived of sucrose. Flies were also retested using the capillary tube method. Mice that were fed upon or probed by the flies were monitored for evidence of exposure to E. risticii. Mice were bled weekly for 6 wk from the tail and the plasma frozen at — 20°C until it was assayed. The plasma was tested for antibody to E. risticii by an indirect immunofluorescence assay (IFA) (Rikihisa et al. 1987). Plasma was initially tested at a 1/20 dilution. Positive and negative controls were included on each slide. Antigen-containing slides were prepared by growing E. risticii in P388-DJ cells, as described above, until the cells were >80% infected. The cells were dislodged with a cell scraper and resuspended by shaking. The suspension was centrifuged at 1,500 x gfor10 min at 4°C. The pellet was washed once in RPMI 1640 medium and resuspended in 0.1 M phosphate-buffered saline (pH 7.3) containing 0.5% (wt/vol) bovine serum albumin (PBS-BSA). The suspension was dotted onto 24-well template slides (Cell-Line Associates, Newfield, NJ), air dried, and fixed in acetone for 10 min. Prepared slides were stored at -20°C until they were tested by IFA. Before testing, antigen-containing slides were washed once with water, the wells covered with 10% (vol/vol) normal goat serum in PBS, placed in a humidified chamber, and incubated for 30 min at 37°C. After a 5-min wash in PBS, the wells were covered with dilutions of the mouse plasma to be tested made in PBS-BSA, and incubated for 30 min at 37°C. Slides were then washed four times (10 min each) in PBS. A 1/50 dilution of FITC-labeled goat-antimouse IgG (H+L) (Cat. no. 1211-0081, Organon Teknika Corp., Durham, NC) in PBS-BSA was applied to each well, incubated for 30 min at 37°C, and washed in PBS four times (10 min each). In the last wash, 0.1-0.2 ml 0.125% (wt/vol) Evans Blue per 20-40 ml of wash was added to counterstain the P388-D! cells. Slides were dried and coverslips affixed to
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
Fly trapping was conducted from May through the first week in October 1988. Female tabanids were collected with canopy traps with black ball decoys (Catts 1970), supplemented with carbon dioxide (dry ice) as an attractant (Everett & Lancaster 1968). Flies were funneled into a screened collection chamber at the apex of the trap. Aerial netting was used to supplement canopy trap collections (Tallamy et al. 1976). Additional specimens were collected as they entered our vehicle. Flies were transported to the laboratory in insulated containers cooled with frozen gel packs and were held at room temperature until used in the feeding trials. All flies collected were identified with dichotomous keys (Pechuman 1973, Drees et al. 1980); identifications were verified by L. L. Pechuman, Cornell University, Ithaca, NY. Voucher specimens were deposited in the Insect Collection of the Department of Entomology, North Carolina State University. Mice. Adult female Balb/C (Charles Rivers Laboratories, Raleigh, NC) mice were acclimated to the laboratory for at least 1 wk before the experiments were begun. The mice were kept in plastic cages in an Illinois-type ventilated chamber at ambient temperature (~25°C). Mice were provided with water and standard rodent chow throughout the study. Inoculum Preparation. Ehrlichia risticii (obtained from Y. Rikihisa of Ohio State University and C. Pretzman, Ohio Department of Health) was propagated in the continuous murine macrophage cell line P388-Di (ATCC TIB 63, American Type Culture Collection, Rockville, MD) as previously described (Pretzman et al. 1987). We previously found the strain to be pathogenic to both mice and horses, with no loss in infectivity due to passage (unpublished data). The P388-D! cell line was propagated in RPMI 1640 medium (Gibco Laboratories, Grand Island, NY) supplemented with 10% fetal bovine serum held at 37°C in an atmosphere of 5% carbon dioxide in air. When a confluent cell layer was attained, the medium was removed and ^ l O 3 E. risticiiinfected P388-D! cells in 1-2 ml of medium were added to the flask. The flasks were incubated for several hours at 35°C with periodic rocking to distribute the infected cell suspension evenly. Additional fresh medium (4.5 ml) was added to each flask, and the cultures were held at 35°C. Infected cells were harvested for preparation of the inoculum when 95% of the cells became infected with E. risticii, as determined by cytocentrifugation of the nonadherent cells and staining with a triarylmethane—xanthene—thiazine stain (Diff-Quik, American Scientific Products, McGraw Park, IL). The confluent cells were disturbed with a cell scraper and resuspended by shaking. The cells were centrifuged at 1,500 x g for 10 min at 4°C, and the pellet was washed once in RPMI medium. The final pellet was re-
807
808
Vol. 29, no. 5
JOURNAL OF MEDICAL ENTOMOLOGY
the slides with PBS-glycerol mounting medium. The preparation was examined at 250 x under epifluorescence microscopy for specific ehrlichial fluorescence. Feeding on Medium in Capillary Tubes.
Genus Tabanus
Genus Chrypsops
T. americanus Forster T. atratus atratus F. T. fulvulus Wiedemann T. gladiator Stone T. imitans Walker T. lineola F. T. melanocerus Weidemann T. molestus molestus Say T. nigrescens Palisot de Beauvois T. nigripes Wiedemann T. pallidescens Philip T. petiolatus Hine T. quinquevittatus Wiedemann T. sparus milleri Whitney T. subsimilis Bellardi T. sulcifrons Macquart T. trimaculatus Palisot de Beauvois
C. callidus Osten Sacken C. celatus Pechuman C. dacne Philip C. flavidus Wiedemann C. geminatus Wiedemann C. macquarti Philip C. montanus Osten Sacken C. pikei Whitney C. reicherti Fairchild C. univittatus Macquart C. upsilon Philip C. vittatus Wiedemann
medium. The second was then removed and tapped onto teflon template-printed slides, air dried, and fixed in acetone for 10 min. Slides were examined as previously described. Results
Tabanids representing two genera and 29 species were collected (Table 1). Although 83 tabanids were tested by feeding on E. risticiiinfected mice (Table 2), only T. fulvulus and T. pallidescens fed upon the mice. Of the 13 flies that probed infected mice, seven ingested a partial or full blood meal. All 13 flies were transferred to uninfected mice, and 7 took partial or full blood meals. Three of the flies that obtained a partial meal on the first host did not attempt to feed on the second host. Three flies that probed but did not ingest blood from the first host fed to repletion on the second mouse. During the 6-wk monitoring period following fly feeding, no antibody to E. risticii was detected in the plasma of the second group of mice, which were probed by tabanids that had previously probed on infected mice. Flies readily fed on medium in capillary tubes, ingesting from 0.5 to nearly 30 fi\ of medium per fly (Table 3). The amount ingested varied with the size of the fly, but even small flies were capable of taking in relatively large volumes of fluid. Intermittent contractions of the abdomen were evident throughout the feeding trials, and, occasionally, a droplet of clearfluidwas expelled from the anus of the fly. Standard capillary tubes worked well with a wide range of flies; for those with large mouthparts, larger tubes were necessary. The ability of 8 species of Chrysops and 12 species of Tabanus to retain E. risticii on their mouthparts was tested by the capillary tube
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
Horse flies and deer flies were offered infectious meals in a manner similar to that previously described for ixodid ticks (Burgdorfer 1957). Each fly was placed on its dorsal surface and held down by strands of clay. An "arm" of clay supported the capillary tube around the mouthparts of the flies. Standard plain (nonheparinized) glass capillary tubes (1.1-1.2 mm inside diameter) or larger laboratory-prepared tubes (1.5 to 1.8 mm inside diameter) were dipped into a suspension of 1056 10 /ml E. risticii-inkcted P388-Di cells in RPMI 1640 medium containing 10% FBS. The tube was placed over the mouthparts of each fly and supported by the clay arm. Flies were allowed to ingest the suspension for 5 min, the tube was removed, and the flies were held without feeding for 2-5 min. A tube of RPMI 1640 with 10% FBS (no cells) was then lowered over the mouthparts, and the flies were allowed to ingest fluid from the second tube for another 5 min. The volumes offluidingested were recorded for each tube. The contents of the second feeding tubes were initially examined by cytocentrifugation and Diff-Quik (American Scientific Products, McGraw Park, IL) staining. This technique proved unsuitable, and any transferred cells were subsequently detected by an IFA using hyperimmune mouse anti-E. risticii serum. The contents of both tubes were dotted onto the wells of a 30-well printed glass slide, air-dried, and acetone-fixed to the slide. The assay was conducted as previously described above, except that the test serum was replaced with a 1/100 dilution of hyperimmune mouse serum, which served as the antigen detection probe. Hyperimmune serum (IFA titer of 1/2,560) was collected from mice experimentally infected with E. risticii. The entire area of each well (2 mm diameter) was searched at 250 x with epifluorescence microscopy for specific fluorescence of organisms within the cell cytoplasm. Fluorescent particles not associated with cells were not considered ehrlichial organisms for this assay. Positive and negative controls were included in each batch of slides tested. The ability to transfer cells on the surface of a needle was also examined using the same protocol used to evaluate transfer of E. risticii between capillary tubes by flies. Hypodermic needles (22 gauge) were cut to leave a 5-mm shaft. Cell suspensions containing E. risticii in capillary tubes were applied to the needles. The capillary tubes containing marked cells were replaced with a capillary tube containing cell-free
Table 1. Tabanids collected at seven sites in Wake (n = 5), Johnston (n = 1), and Duplin (n = 1) counties, N.C., during early May—October 1988
September 1992
LEVINE ET AL.: MECHANICAL TRANSFER OF
E. risticii BY
809
TABANIDS
Table 2. Exposure time, probing attempts, and engorgement success of T. fulvulus (Tf) and T. pallidescens placed on £ . rwtieii-infected mice and transferred to uninfected mice. Infected mouse Fly no"
Time on host, min
Probe
Tf-NN Tp-5 Tf-19 Tf-20 Tf-21 Tf-22 Tf-24 Tf-102 Tf-116 Tf-175 Tf-208 Tf-209 Tf-213
2 14 15 7 13 10 8 18 4 18 11 5 9
Y Y Y Y Y Y Y Y Y Y Y Y Y
6
Engorgement levelc
period, min
Time on host, min
P P O O P
0 6 1 4 0 3 2 4 60 0.3 1 0.3 0.2
8 22 13 18 10-15 10-15 10-15 6 8 10 3 15 9
o op o oF P P
(Tp)
Uninfected mouse Engorgement Probe6 levelc N Y Y Y N N N Y Y N N Y Y
O F F F O O O P F O O P P
method (Table 3). Ten of 109 (9.2%) flies retained cells on their mouthparts after feeding on medium containing infected cells and transferred the cells to medium in a second tube. No E. risticii were observed in the cells transferred by two flies. Cell transfer was observed in 10 of 10 trials when needles were used as substrate. Discussion Demonstration of the competence of arthropods as vectors requires recognition of infection
in a host after transfer of the agent by the arthropod. Flies that probed on £. mticii-infected mice apparently failed to transfer an infectious dose of the agent to noninfected mice. Previous to this study, we confirmed the validity of the IFA procedure for detection of antibody to E. risticii in murine serum (unpublished data), and IFA provided a viable means of assessing the exposure of mice to E. risticii. Although it is unlikely that mouse-to-mouse transmission by flies occurred, a small number of infected cells may have been transferred but were insufficient
Table 3. Mean volume (±SD) of infected cell suspension and medium ingested in 5 min by tabanids individually fed by the capillary tube method, and transfer of cells as determined by immunofluorescence assay Species Chrysops spp. C. callidus C. celatus C. dacne C.flavidus C. geminatus C. montanus C. univittatus C. vittatus
C. spp.
Tabanusspp. T. americanus T. atratus atratus T. fuloulus T. lineola T. melanocerus T. nigrescens T. nigripes T. petiolatus T. quinquevittatus T. sparus milleri T. subsimilis T. sulcifrons T. spp. Undetermined tabanids Total
No.
2 5 2 5 1 1 1 30 33 3 1 1 1 3 1 1 1 3 9 3 17 3 7 134
° ND, not done. b Larger laboratory-prepared tubes were used.
Vol cell suspension, fi\ 1.0 3.3 2.0 4.6 1.0 1.0 3.0 2.5 1.7
(2.6) (5.7)
(2.1) (1.5)
9.8 (5.0) 28.0b 3.5 16.0 6.0 (6.2) 1.0 6.0 2.0 2.3 (1.2) 3.4 (2.8) 1.7 (1.2) 7.2 (5.7) 8.2 (6.9) 7.5 (9.2)
Vol medium, /xl 3.0 1.4 0.8 3.3 1.0 0.5 0 3.3 1.6
(1.1) (6.0)
(2.9) (1.3)
3.0 (2.5) 12.4fc 4.0 5.0 1.8 (0.8) 1.0 2.0 1.5 1.0 (0.5) 2.6 (1.5) 0.8 (0.3) 11.0 (13.3) 3.8 (1.3) 2.9 (1.9)
No. positive/ total tested" 0/ 2 0/ 4 ND 0/ 4 ND 0/ 1 0/ 1 1/26 (3.8%) 3/25 (12%) 1/ 3 (33.3%) 0/ 1 ND 0/ 0 0/ 3 0/ 1 0/ 1 0/ 1 0/ 3 0/ 7 0/ 3 4/14 (28.6%) 0/ 3 1/ 6 (16.7%) 10/109 (9.2%)
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
" Tf, Tabanus fulvulus; Tp, Tabanus pallidescens. b Probe success: Y, yes; N, no. c Engorgement level: O, no visible abdomen expansion; P, partial blood meal taken; F, full blood meal taken.
810
JOURNAL OF MEDICAL ENTOMOLOGY
of infected cells that can potentially be ingested or contaminate the mouthparts of a fly during feeding. Actual volumes of blood and tissue fluid ingested, however, will vary with the species of fly feeding. Foil et al. (1989), in studies conducted with T.fusciostatus and bovine leukemia virus, found that as few as 10fliestransferred that virus from infected to noninfected hosts. The authors estimated that 10flieswould move up to 60 /Ltl of infected blood between hosts. The actual potential for transmission is also dependent on the number of infected cells present at the time of feeding (Foil et al. 1989). Precise verification of the ehrlichemia developed by infected horses and the actual number of E. risticii cells retained in bloodmeal residue on fly mouthparts is needed to quantify the potential for tabanid transmission accurately. Mechanical transmission occurs when a fly feeding on a host with a patent infection is interrupted and the arthropod moves to a susceptible host. The vector potential of each fly species for a given pathogen, however, is quite variable and is dependent on: (1) the stability of the agent on fly mouthparts, (2) the quantity of infected material that can be transferred between hosts, and (3) the abundance and biting habits of individual fly species (Foil 1989). Host skin thickness, the response of the host to fly biting, and the maintenance of a patent infection determine the hosts' ability to support mechanical transmission by a competent vector (Foil et al. 1985, Foil 1989). The spatial and temporal distribution of immune and susceptible hosts also affect the potential for mechanical transfer of an agent. Epidemiologic studies at locations where equine monocytic ehrlichiosis has been diagnosed suggest that cases are generally spatially isolated (Palmer et al. 1986, Burg et al. 1990) and that mechanical transmission of E. risticii between horses might be unlikely. However, in one study, antibody to E. risticii was detected in the serum of cats residing on farms where cases of EME had been observed (Perry et al. 1989). Horses may be dead-end hosts (Burg et al. 1990), and the role of cats or other hosts as potential reservoirs that help support E. risticii transmission may warrant additional study (Perry et al. 1989). Our studies demonstrate that E. risticiiinfected cells can be retained on the mouthparts of some species of tabanids. However, additional experiments with infected and susceptible horses and possibly other hosts are needed to document the vector-competence of tabanids and the capacity of individual species to maintain transmission after the pathogen is introduced into an equine population. Acknowledgment We thank L. L. Pechuman (Cornell University) for confirming the identity of the tabanids, D. Moncol and
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
to induce an antibody response. Isolation of E. risticii would have provided a direct alternative measure of infection, but it was not attempted. Three species of tabanids fed by the capillary tube method retained cells on their mouthparts and transferred infected cells between the capillary tubes, demonstrating that E. risticiiinfected cells can be successfully carried on the mouthparts of tabanids. Studies conducted with T.fuscicostatus have shown that ^lO nl of residual blood remain on the mouthparts of this species when feeding on a host is interrupted (Foil et al. 1987). The period of time that an agent remains viable on the mouthparts offliesis quite variable and is dependent on related factors (pathogen, environmental, vector) (Foil 1989). We used an interfeed period of 2—5 min between capillary tubes, and E. m£icu-infected cells remained intact on the mouthparts of at least three species during this period. Bloodmeal residue has been detected on tabanid mouthparts hours after the interruption of feeding, but detectable residues decline significantly after several hours (Foil et al. 1987). Fly grooming and mouthpart cleaning may account for some reduction in residues. Tabanids are recognized mechanical vectors of >30 pathogens that cause disease in animals (Krinsky 1976). Tabanid biting activity can be intense. In some areas, tabanids have been observed landing on horses at a rate of 1,000 flies per hour. Several species of tabanids may serve as mechanical vectors, and transfer of an infective dose may occur through the combined attack of multiple species (Foil 1989). Actual biting rates in a pasture will vary with the species of flies present, the presence of suitable habitat, climate, herd size, animal behavior, and the distribution of animals. Although the ability of numerous species of biting flies to transmit pathogens has been well documented, no Ehrlichia sp. is known to be fly-transmitted. The Ehrlichieae are obligate intracellular parasites (Ristic & Huxsoll 1984), and successful mechanical transmission requires the transfer of intact infected cells between hosts. Tabanids and iatrogenic spread by needle are recognized means of transmission of other cellassociated pathogens such as equine infectious anemia virus (Hawkins et al. 1976, Issel et al. 1988), bovine leukemia virus (Bech-Nielson et al. 1978, Foil et al. 1988), and Anaplasma marginale (Dikmans 1950). These studies suggest that E. risticii cells may remain viable on the mouthparts of some species of tabanids and on the surface of needles. Biologic transmission of E. risticii by Stomoxys calcitrans has also been evaluated. However, this species failed to transfer infection from an infected to a noninfected host (Burg et al. 1990). The concentration of a pathogen in the blood and local tissues of a host determine the number
Vol. 29, no. 5
September 1992
LEVINE ET AL.: MECHANICAL TRANSFER OF E. risticii BY TABANIDS
A. Artis for technical assistance, and E. W. Cupp (University of Arizona) for suggesting the tube-feeding method. Funds to support this research were provided by the Morris Animal Foundation.
References Cited
investigation of farms with Potomac horse fever (equine monocytic ehrlichiosis). Acta Vet. Scand. (Suppl.) 84: 319-322. Hahn, N. E., M. Fletcher, R. M. Rice, K. M. Kocan, J. W. Hansen, J. A. Hair, R. W. Barker & B. D. Perry. 1990. Attempted transmission of Ehrlichia risticii, causative agent of Potomac horse fever, by the ticks, Dermacentor variabilis, Rhipicephalus sanguineus, Ixodes scapularis, and Amblyomma americanum. Exp. Appl. Acarol. 8: 41-50. Hawkins, J. A., W. V. Adams, Jr., B. H. Wilson, C. J. Issel & E. E. Robl. 1976. Transmission of equine infectious anemia virus by Tabanus fuscicostatus. J. Am. Vet. Med. Assoc. 168: 63-64. Holland, C. J., E. Weiss, W. Burgdorfer, A. I. Cole & I. Kakoma. 1985a. Ehrlichia risticii sp. nov.: etiological agent of equine monocytic ehrlichiosis (synonym, Potomac horse fever). Int. J. Syst. Bacteriol. 35: 524-526. Holland, C. J., M. Ristic, A. I. Cole, P. Johnson, G. Baker & T. Goetz. 1985b. Isolation, experimental transmission, and characterization of causative agent of Potomac horse fever. Science 227: 522524. Issel, C. J., K. Rushlow, L. D. Foil & R. C. Montelaro. 1988. A perspective on equine infectious anemia with an emphasis on vector transmission and genetic analysis. Vet. Microbiol. 17: 251-286. Knowles, R. C , D. W. Anderson, W. D. Shipley, R. H. Whitlock, B. D. Perry & J. P. Davidson. 1984. Acute equine diarrhea syndrome (AEDS): a preliminary report. Proc. Am. Assoc. Equine Pract. 29: 353-357. Krinsky, W. L. 1976. Animal disease agents transmitted by horse flies and deer flies (Diptera: Tabanidae). J. Med. Entomol. 13: 225-275. Levine, J. F., M. G. Levy, W. L. Nicholson & R. B. Gager. 1990. Attempted Ehrlichia risticii transmission with Dermacentor variabilis (Acari: Ixodidae). J. Med. Entomol. 27: 931-933. Palmer, J. E., R. H. Whitlock & C. E. Benson. 1986. Equine ehrlichial colitis (Potomac horse fever): recognition of the disease in Pennsylvania, New Jersey, New York, Ohio, Idaho, and Connecticut. J. Am. Vet. Med. Assoc. 189: 197-199. Pechuman, L. L. 1973. Horse flies and deer flies of Virginia (Diptera: Tabanidae). Va. Polytech. Inst. State Univ. Res. Div. Bull. 81. Perry, B. D., Y. Rikihisa & G. K. Saunders. 1985. Intradermal transmission of Potomac horse fever. Vet. Rec. 16: 246-274. Perry, B. D., E. T. Schmidtmann, R. M. Rice, J. W. Hansen, M. Fletcher, E. C. Turner, M. G. Robl & N. E. Hahn. 1989. Epidemiology of Potomac horse fever: an investigation into the possible role of nonequine mammals. Vet. Rec. 125: 83-86. Pretzman, C. I., Y. Rikihisa, D. Ralph, J. C. Gordon & S. Bech-Nielson. 1987. Enzyme-linked immunosorbent assay for Potomac horse fever disease. J. Clin. Microbiol. 25: 31-36. Rikihisa, Y. & B. D. Perry. 1985. Causative ehrlichial organisms in Potomac horse fever. Infect. Immun. 49: 513-517. Rikihisa, Y., G. C. Johnson & C. J. Burger. 1987. Reduced immune responsiveness and lymphoid depletion in mice infected with Ehrlichia risticii. Infect. Immun. 55: 2215-2222. Ristic, M. & D. L. Huxsoll. 1984. Tribe II. Ehr-
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
Bech-Nielsen, S., C. E. Piper & J. F. Ferrer. 1978. Natural mode of transmission of the bovine leukemia virus: role of bloodsucking insects. Am. J. Vet. Res. 39: 1089-1092. Burg, J. G., A. W. Roberts, N. M. Williams, D. G. Powell & F. W. Knapp. 1990. Attempted transmission of Ehrlichia risticii (Rickettsiaceae) with Stomoxys calcitrans (Diptera: Muscidae). J. Med. Entomol. 27: 874-877. Burg, J. G., D. G. Powell & F. W. Knapp. 1991. Arthropod faunal composition on Kentucky equine premises. J. Med. Entomol. 28: 658-662. Burgdorfer, W. 1957. Artificial feeding of ixodid ticks for studies on the transmission of disease agents. J. Infect. Dis. 100: 212-214. Carroll, J. F. & E. T. Schmidtmann. 1986. American dog tick (Acari: Ixodidae) summer activity on equine premises enzootic for Potomac horse fever in south-central Maryland. J. Econ. Entomol. 79: 62-66. Catts, E. P. 1970. A canopy trap for collecting Tabanidae. Mosq. News 30: 472-474. Dikmans, G. 1950. The transmission of anaplasmosis. Am. J. Vet. Res. 11: 5-16. Drees, B. M., L. Butler & L. L. Pechuman. 1980. Horse flies and deer flies of West Virginia: an illustrated key (Diptera, Tabanidae). W. Va. Univ. Agric. For. Exp. Stn. Bull. 674. Dutta, S. K., A. C. Myrup, R. M. Rice, M. G. Robl & R. C. Hammond. 1985. Experimental reproduction of Potomac horse fever in horses with a newly isolated Ehrlichia organism. J. Clin. Microbiol. 22: 265-269. Everett, R. & J. L. Lancaster. 1968. A comparison of animal and dry-ice baited traps for the collection of tabanids. J. Econ. Entomol. 61: 863-864. Fletcher, M. G., E. C. Turner & J. W. Hansen. 1988. Horse-baited insect trap and mobile insect sorting table used in a disease vector identification study. J. Am. Mosq. Control Assoc. 4: 431-435. Foil, L. D. 1989. Tabanids as vectors of disease agents. Parasitol. Today 5: 88-96. Foil, L. D., D. Stage, W. V. Adams, Jr. & C. J. Issel. 1985. Observations of tabanid feeding on mares and foals. Am. J. Vet. Res. 46: 1111-1113. Foil, L. D., W. V. Adams, J. M. McManus & C. J. Issel. 1987. Bloodmeal residue on mouthparts of Tabanus fuscicostatus (Diptera: Tabanidae) and the potential for mechanical transmission of pathogens. J. Med. Entomol. 24: 613-616. Foil, L. D., C. L. Seger, D. D. French, C. J. Issel, J. M. McManus, C. L. Ohrberg & R. T. Ramsey. 1988. Mechanical transmission of bovine leukemia virus by horse flies (Diptera: Tabanidae). J. Med. Entomol. 25: 374-376. Foil, L. D., D. D. French, P. G. Hoyt, C. J. Issel, D. J. Leprince, J. M. McManus & C. L. Seger. 1989. Transmission of bovine leukemia virus by Tabanus fuscicostatus. Am. J. Vet. Res. 50: 1771-1773. Gordon, J. C , S. Bech-Nielson, C. Kohn, W. Farrar, M. Parsons & W. Foster. 1988. An epidemiological
811
812
JOURNAL O F MEDICAL ENTOMOLOGY
lichieae, pp. 704-709. In N. R. Krieg [ed.], Bergey's manual of systematic bacteriology, vol. 1. Williams & Wilkms, Baltimore. Schmidtmann, E. T., M. G. Robl & J. F. Carroll. 1986. Attempted transmission of Ehrlichia risticii by field-captured Dermacentor variabilis (Acari: Ixodidae). Am. J. Vet. Res. 47: 2393-2395.
Vol. 29, no.
5
Tallamy, D. W., E. J. Hansens & R. F. Denno. 1976. A comparison of Malaise trapping and aerial netting for sampling a horse fly and deer fly community. Environ. Entomol. 5: 788-792. Received for publication 2 December 1991; accepted 6 April 1992.
Downloaded from http://jme.oxfordjournals.org/ by guest on January 13, 2016
