JOURNAL OF CLINICAL MICROBIOLOGY, July 1999, p. 2262–2269 0095-1137/99/$04.0010 Copyright © 1999, American Society for Microbiology. All Rights Reserved.
Vol. 37, No. 7
Upper Respiratory Tract Disease in the Gopher Tortoise Is Caused by Mycoplasma agassizii† M. B. BROWN,1* G. S. MCLAUGHLIN,2,3 P. A. KLEIN,4 B. C. CRENSHAW,1 I. M. SCHUMACHER,4 D. R. BROWN,1 AND E. R. JACOBSON2 Department of Pathobiology and Division of Comparative Medicine1 and Department of Small Animal Clinical Sciences,2 College of Veterinary Medicine, Department of Wildlife Ecology and Conservation, Institute of Food and Agricultural Sciences,3 and Department of Pathology, Immunology, and Laboratory Medicine, College of Medicine,4 University of Florida, Gainesville, Florida Received 19 November 1998/Returned for modification 18 February 1999/Accepted 31 March 1999
Upper respiratory tract disease (URTD) has been observed in a number of tortoise species, including the desert tortoise (Gopherus agassizii) and the gopher tortoise (Gopherus polyphemus). Clinical signs of URTD in gopher tortoises are similar to those in desert tortoises and include serous, mucoid, or purulent discharge from the nares, excessive tearing to purulent ocular discharge, conjunctivitis, and edema of the eyelids and ocular glands. The objectives of the present study were to determine if Mycoplasma agassizii was an etiologic agent of URTD in the gopher tortoise and to determine the clinical course of the experimental infection in a dose-response infection study. Tortoises were inoculated intranasally with 0.5 ml (0.25 ml/nostril) of either sterile SP4 broth (control group; n 5 10) or 108 color-changing units (CCU) (total dose) of M. agassizii 723 (experimental infection group; n 5 9). M. agassizii caused clinical signs compatible with those observed in tortoises with natural infections. Clinical signs of URTD were evident in seven of nine experimentally infected tortoises by 4 weeks postinfection (p.i.) and in eight of nine experimentally infected tortoises by 8 weeks p.i. In the dose-response experiments, tortoises were inoculated intranasally with a low (101 CCU; n 5 6), medium (103 CCU; n 5 6), or high (105 CCU; n 5 5) dose of M. agassizii 723 or with sterile SP4 broth (n 5 10). At all time points p.i. in both experiments, M. agassizii could be isolated from the nares of at least 50% of the tortoises. All of the experimentally infected tortoises seroconverted, and levels of antibody were statistically higher in infected animals than in control animals for all time points of >4 weeks p.i. (P < 0.0001). Control tortoises in both experiments did not show clinical signs, did not seroconvert, and did not have detectable M. agassizii by either culture or PCR at any point in the study. Histological lesions were compatible with those observed in tortoises with natural infections. The numbers of M. agassizii 723 did not influence the clinical expression of URTD or the antibody response, suggesting that the strain chosen for these studies was highly virulent. On the basis of the results of the transmission studies, we conclude that M. agassizii is an etiologic agent of URTD in the gopher tortoise. Gopher tortoises (Gopherus polyphemus) are found in the southeastern United States, with the major population concentrations found in Florida and southern Alabama and Georgia and only remnant populations found in South Carolina, Mississippi, and Louisiana (9). The gopher tortoise is legally protected in all states within the range (Alabama, Mississippi, Louisiana, Georgia, South Carolina, and Florida) and is listed in Appendix II of the Convention on International Trade in Endangered Species of Wild Fauna and Flora, which requires permits for the exportation of the species from the United States to any signatory nation or for reexportation (20). Gopher tortoises are an important element in the ecosystems in which they are found and are considered by many ecologists to be a keystone species. Gophers are the most fossorial of the four North American species of tortoises, digging burrows that may extend 5 m down from the surface and 15 m in length (9, 12). The burrows provide a microclimatically stable environment not only for the tortoises but also for numerous commensal species. Approximately 60 vertebrate species, from snakes to birds, and over 300 invertebrates including spiders,
crickets, and beetles have been found in tortoise burrows or have been observed using them as permanent homes or refuges from heat, cold, fire, and predators (14, 22, 36). Several species that either exclusively or frequently use tortoise burrows have legal protection in Florida and other parts of their ranges (6). Thus, tortoises are of critical importance to the ecosystem. Upper respiratory tract disease (URTD) has been observed in a number of tortoise species (15, 16, 19), including the desert tortoise (Gopherus agassizii) and the gopher tortoise. Clinical signs of URTD have been observed in a number of imported captive tortoise species (19) and in tortoises submitted to the Veterinary Medical Teaching Hospital (VMTH), University of Florida (UF), including the red-footed tortoise (Geochelone carbonaria), leopard tortoise (Geochelone pardalis), Indian star tortoise (Geochelone elegans), and radiated tortoise (Geochelone radiata). Numerous wild and captive gopher tortoises have been submitted to VMTH with clinical signs consistent with URTD. Clinical signs of URTD in gopher and desert tortoises are similar and include serous, mucoid, or purulent discharge from the nares, excessive tearing to purulent ocular discharge, conjunctivitis, and edema of the eyelids and ocular glands (16, 31). Individual infected tortoises vary in the suite of signs that they have, and the severity can vary from day to day. Nares may become occluded with caseous exudate, preventing externally visible nasal discharge. Tortoises may become lethargic and
* Corresponding author. Mailing address: Department of Pathobiology, Box 110880, College of Veterinary Medicine, University of Florida, Gainesville, FL 32611-0880. Phone: (352) 392-4700, ext. 3970. Fax: (352) 846-2781. E-mail:
[email protected]. † Journal series article R-06916 of the Florida Agriculture Experiment Station. 2262
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anorectic, leading to dehydration, emaciation, and eventual death from cachexia. In a previous study, we fulfilled Koch’s postulates and demonstrated that an etiologic agent for URTD in the desert tortoise is Mycoplasma agassizii proposed sp. novum (2, 4). Histologically, the lesions from experimentally infected desert tortoises were consistent with those seen in naturally infected tortoises (4, 16). In the desert tortoise, we have shown that the presence of clinical signs of URTD was positively related to the presence of specific antibody to M. agassizii (31). Additional work led to the development of a PCR test for detection of the bacteria in nasal lavage and swab samples (2). We isolated M. agassizii from the nasal passages of clinically ill gopher tortoises submitted to VMTH, UF. The similarity of the clinical signs and histological lesions between experimentally infected and naturally infected tortoises and the isolation of M. agassizii from naturally infected gopher tortoises suggested that URTD in this species might also be of mycoplasmal origin. The objectives of this study were to determine if M. agassizii was an etiologic agent of URTD in the gopher tortoise and to determine the clinical course of the experimental infection in a dose-response infection study. MATERIALS AND METHODS Tortoises. Gopher tortoises were transferred under Florida Game and Fresh Water Fish Commission permit nos. WX93227, issued to E. R. Jacobson, and WX94037, issued to M. B. Brown, from a development site in central Florida in April, July, and August 1994 and April 1995 and were processed on the day following arrival. Tortoises were examined for clinical signs of URTD: nasal and ocular discharge, palpebral edema, and conjunctivitis. Tortoises were weighed to the nearest 10 g, and ketamine hydrochloride (Ketaset; Fort Dodge Laboratories, Inc., Fort Dodge, Iowa) was administered at 20 mg/kg of body weight. A blood sample (2 to 3 ml) was drawn from the jugular or brachial vein and was placed in a Vacutainer tube (Becton Dickinson and Company, Rutherford, N.J.) containing lithium heparin. Blood was centrifuged, and an aliquot of plasma was removed for specific antibody screening by an enzyme-linked immunosorbent assay (ELISA). After cleansing of the area around the nares with alcoholdampened gauze, nasal lavage samples were collected by flushing with approximately 0.5 ml of sterile SP4 broth with a 1-ml syringe without a needle. Calcium alginate-tipped swabs were gently inserted into the nares, and a sample was obtained and streaked onto SP4 agar plates (33). Husbandry. Tortoises were housed individually in outdoor pens at the UF Animal Resource Farm. There were four groups of 10 pens in a larger enclosure surrounded by a chain-link fence. Individual pens were approximately 21 m2, were constructed of a wooden frame with sheet metal extending vertically approximately 0.7 m above and below the ground, and were partially covered by shade cloth. The tortoises were provided an artificial burrow, a water dish, and a cement feeding stone. Because tortoises burrow, the risk of cross contamination was too great to allow randomization of treatment groups within pen groups. Each treatment group of 10 pens was separated from the other treatment pens by .200 m. The tortoises were fed a salad of mixed vegetables three times per week, and fruit was provided on an occasional basis. Water was provided as needed. Husbandry personnel wore gloves for all procedures requiring handling of food, feeding stones, or water dishes. Entry into pens and handling of tortoises were restricted to research personnel. Any person handling a tortoise wore clean gloves, which were changed as necessary and before handling of a different tortoise. Infection groups. Animals were allowed to acclimate for a minimum of 1 month prior to initiation of the experimental transmission trials. Tortoises were blocked on the basis of size and sex prior to random assignment to experimental and control groups. Tortoises in the control group (n 5 10) were sham inoculated intranasally with 0.5 ml (0.25 ml/nostril) of sterile SP4 broth. Tortoises in the experimental infection group (n 5 9) were inoculated intranasally with 0.5 ml (0.25 ml/nostril; 108 color-changing units [CCU] [total dose]) of M. agassizii 723. Isolate 723 was obtained from a clinically ill tortoise from Sanibel Island, Lee County, Fla. M. agassizii 723 was grown in SP4 broth and was only two passages from the primary isolation. Aliquots of strain 723 were frozen at 280°C, and all infections were done with a common stock of M. agassizii. The purity of the isolate was determined by growth inhibition and 16S rRNA sequence analysis (2). Dose-response study. After the initial experiments which demonstrated that M. agassizii caused URTD in the gopher tortoise, an experiment was designed to determine the effect of dose on clinical expression of disease and immune response. Tortoises were inoculated intranasally with a low (101 CCU; n 5 6), medium (103 CCU; n 5 6), or high (105 CCU; n 5 5) dose of M. agassizii. Tortoises in the control group (n 5 10) were sham inoculated intranasally with
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0.5 ml (0.25 ml/nostril) of sterile SP4 broth. Monitoring and housing were identical to those in the initial experimental infection study. Postinfection monitoring of tortoises. The tortoises were observed from 3 to 7 days each week, with some days including multiple observations. Because of individual behavior patterns, not every tortoise was observed at each daily observation time point. At specific time points (usually 2- to 4-week intervals, depending on the study), tortoises were captured by hand or with wire cage-type traps (Tomahawk Live Trap Company, Tomahawk, Wis.) that were covered with brown paper to protect the animals from the weather. The traps were cleaned, sprayed with bleach solution, and allowed to air dry following each use. The paper was discarded, and fresh paper was used for the next trapping effort. Each tortoise was placed in a plastic, lidded container (LEWISystems; Menasha Corporation, Watertown, Wis.) for transport and holding. Containers were bleached, scrubbed, and washed in an automatic cage washer before reuse. Tortoises were examined for clinical signs of URTD: nasal and ocular discharge, palpebral edema, and conjunctivitis. A photographic record consisting of right, left, and full face views was made for each tortoise at each time point when the tortoises were captured. The signs were graded individually on a scale of from 0 to 3, which indicated none, minimal, mild, and severe signs, respectively. Visual grading of signs was confirmed by independent observation of the photographic record. Serum was obtained for quantitation of specific antibody. Nasal swabs and lavages were obtained for culture and PCR testing. Culture. Mycoplasmal cultures were performed as described previously (4). A 100-ml aliquot of the lavage sample was used for PCR analysis; the remaining sample was serially diluted 10-fold to 1022 and was incubated at 30°C for a maximum of 3 weeks or until it was determined to be positive or contaminated. In some cases, an aliquot of the broth culture of both lavages and swabs was removed after 24 to 48 h and was used for PCR to confirm growth of M. agassizii. Twenty microliters of each dilution was placed on SP4 agar, and the plates were incubated at 30°C in 5% CO2. The swabs were streaked onto the surface of an SP4 plate. The plates were examined regularly for a maximum of 6 weeks to detect the growth of mycoplasma. PCR. Nasal aspirate lavage specimens were analyzed for the presence of M. agassizii DNA on the basis of PCR amplification of the 16S rRNA gene (2). Nasal lavage specimens and selected culture samples obtained at between 24 and 48 h were centrifuged at 16,000 3 g for 60 min at 4°C, and the supernatant was aspirated. The pellets were resuspended in 3 to 4 ml of 20 mg of proteinase K (Sigma, St. Louis, Mo.) per ml in 20 ml of lysis buffer (100 mM Tris [pH 7.5], 6.5 mM dithiothreitol, 0.05% Tween 20), and the mixture was incubated at 37°C for 8 to 16 h. After denaturation of the proteinase K at 97°C for 15 min, 5 ml of each sample was removed and was added to 45 ml of a reaction solution containing two primers for the 16S rRNA gene at 1 mM each, deoxynucleoside triphosphates at 200 mM, 2.0 mM MgCl2, and 2.5 U of Taq polymerase (Promega, Madison, Wis.). The primers were complementary to sequences found in the V3 variable region of the 16S rRNA gene (sense strand nucleotides [nt] 471 to 490 [59-CC TATATTATGACGGTACTG-39]) and a Mycoplasma genus-specific region (anti-sense strand nt 1055 to 1031 [59-TGCACCATCTGTCACTCTGTTAACCTC39]) (2, 34). The samples were subjected to 50 cycles of template denaturation for 45 s at 94°C, primer annealing for 1 min at 55°C, and polymerization for 45 s at 72°C, followed by 10 min at 72°C. Positive samples yielded 576-bp products that were visualized by combining 15 ml of product with 2 ml of bromphenol blue in 50% glycerol solution and electrophoresing on ethidium bromide-stained 1.5% agarose gels in Tris-borate-EDTA buffer. Positive control samples with 250 ng of purified M. agassizii DNA as the template and negative control samples with water in place of a template were included with each amplification run. A molecular weight marker, an HaeIII digest of phage fX174 DNA, was included on each gel. Restriction fragment length polymorphism analysis was conducted with at least one isolate from each tortoise in order to confirm conclusively that the isolates obtained from naturally and experimentally infected tortoises were M. agassizii (2). Twenty-microliter samples of products from the amplification procedure described above were incubated with 10 to 20 U of the endonuclease AgeI (New England Biolabs, Inc., Beverly, Mass.), which cuts the M. agassizii amplification product at nt 613, and 5 ml of reaction buffer at 25°C for 1 h, and the products were electrophoresed as described above. The procedure resulted in products of 434 and 142 bp from M. agassizii-positive samples, and all mycoplasmal isolates in the study were confirmed to be M. agassizii. ELISA. Specific antibody to M. agassizii was determined by ELISA as described previously (30). Ninety-six-well microliter plates (Maxisorp F96; Nunc, Kamstrup, Denmark) were coated with 50 ml of a whole-cell lysate of M. agassizii 723 at 10 mg/ml in phosphate-buffered saline (PBS) with 0.02% azide (PBS-AZ). The plates were incubated overnight at 4°C, washed four times with PBS-AZ plus 0.05% Tween 20 (PBST) in an automatic plate washer (EL403; Bio-Tek Instruments, Inc., Winooski, Vt.), and blocked overnight at 4°C with 250 ml of PBST containing 5% nonfat dry milk (PBS-TM) per well. Following washing, 50 ml of plasma diluted appropriately for the specific study with PBS-TM was added to individual wells in duplicate or triplicate, and the plates were incubated at room temperature for 60 min. The plates were washed, 50 ml (per well) of a biotinylated monoclonal antibody (monoclonal antibody HL673) against the light chain of desert tortoise immunoglobulins Y and M at 1 mg/ml in PBS-TM was added, and the plates were incubated for 60 min. Following washing, a conjugate of alkaline phosphatase and streptavidin (Zymed Laboratories, Inc., San Francisco,
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Calif.) at 1:2,000 in PBS-AZ was added at 50 ml/well, and the plates were incubated for 60 min and washed. The substrate, p-nitrophenyl phosphate disodium (pNPP; Sigma), was prepared at 1 mg/ml in 0.01 M sodium bicarbonate (pH 9.6) with 2 mM MgCl2 and was added to the wells at 100 ml/well. The plates were incubated for 60 min in the dark and were then read at 405 nm on a microplate reader (EAR 400 AT; SLT Labinstruments, Salzburg, Austria). The mean for two or three wells coated with antigen and incubated with conjugate and substrate only was used as the blank. Seroconversion was defined as an antibody level greater than 2 standard deviations above the values for normal control serum. A positive control, which consisted of plasma from a naturally infected gopher tortoise from Sanibel Island, Fla., and a negative control, which consisted of plasma from an uninfected tortoise from Orange County, Fla., were included on each plate. Pathology. After a minimum of 16 weeks postinfection (p.i.), all diseased and selected healthy tortoises were killed with a combination of drugs. Ketamine was administered intramuscularly at 60 to 80 mg/kg followed by administration of a concentrated barbiturate solution (Socumb; The Butler Company, Columbus, Ohio) intracoelomically at 1 ml/kg. Once the tortoises showed complete muscle relaxation and were unresponsive to painful stimulation, they were exsanguinated with a 23-gauge butterfly catheter inserted into the carotid artery and then decapitated. Lavage and swab samples were collected as described previously, and then the head was bisected longitudinally with an electric saw. Following bisection, the cartilage over each nasal cavity was reflected aseptically, and lavage and swab specimens from both left and right nasal cavities were collected. For those tortoises selected for complete necropsy, the plastron was removed from the carapace, and viscera within the coelomic cavity were exposed. A gross necropsy was conducted. For histopathologic studies, the heads were fixed in 10% neutral buffered formalin, decalcified, embedded in paraffin, sectioned longitudinally at 5 to 6 mm, and stained with hematoxylin and eosin. Sections were examined by light microscopy and were classified on a scale of from 0 to 5, with 0 being normal and 5 exhibiting severe inflammation and/or changes. Changes in the epithelium and submucosa were recorded separately. The following criteria were used for the grading of lesions: (i) normal (score 5 0), occasional small subepithelial lymphoid aggregates, rare heterophils in the lamina propria, no changes in mucosal or glandular epithelium, and no edema; (ii) mild (score 5 1), multifocal small subepithelial lymphoid aggregates; multifocally, small numbers of heterophils, lymphocytes, and plasma cells in the lamina propria; mild edema in the lamina propria; and minimal changes in mucosal epithelium; (iii) moderate (score 5 2 or 3), multifocal to focally extensive lymphoid aggregates; diffusely, moderate numbers of heterophils, lymphocytes, and plasma cells in the lamina propria, occasionally infiltrating the overlying mucosal epithelium; moderate edema in the lamina propria; and proliferation and disorganization of the basal epithelium; (iv) severe (score 5 4 or 5), focally extensive to diffuse bands of lymphocytes and plasma cells subjacent to and obscuring the overlying mucosal epithelium; large numbers of heterophils in lamina propria and infiltrating the overlying mucosal epithelium; marked edema of the lamina propria; degeneration, necrosis, and loss of the mucosal epithelium with occasional erosion; proliferation of the basal cells of the epithelium with metaplasia of the mucous and olfactory epithelium to a basaloid epithelium; and occasional squamous metaplasia. Statistical analysis. Binomial data were analyzed by the chi-square test. Continuous data were analyzed by t test (two-group comparisons) or analysis of variance (multiple comparisons), followed by Fisher’s least-squares-difference analysis. For clinical sign score data, the nonparametric Mann-Whitney (twogroup comparison) or the Kruskal-Wallis (comparison of more than two groups) test was used. A P value of ,0.05 was accepted as significant.
RESULTS Clinical disease outcome: experimentally infected tortoises. M. agassizii caused clinical signs compatible with those observed in animals with natural infection (Table 1; see also Fig. 2). Clinical signs of URTD were evident in seven of nine experimentally infected tortoises by 4 weeks p.i. and in eight of nine experimentally infected tortoises by 8 weeks p.i. (Table 1). The one tortoise which failed to show clinical signs did seroconvert, indicating that the animal had been colonized sufficiently to stimulate a host immune response. At all time points p.i., M. agassizii could be isolated from the nares of at least 50% of the tortoises (Table 1). After 8 weeks p.i., the ELISA was the most reliable method of detection, with 100% of the infected tortoises testing positive for antibodies to M. agassizii (Table 1). Control tortoises which were sham inoculated with sterile broth did not show clinical signs and did not seroconvert, and M. agassizii was not detected in these tortoises by either culture or PCR at any point in the study. The cumulative scores for infected and control tortoises
TABLE 1. Outcome of transmission study to determine pathogenicity of M. agassizii in gopher tortoisesa No. of animals positive for the following/ total no. of animals tested (%):
Wk
0 4 8 12 16
Clinical signs
ELISA
PCR
Culture
0/9 (0) 7/9 (77.8) 8/9 (88.9) 8/9 (88.9) 8/9 (88.9)
0/9 (0) 3/6 (33.3) 8/9 (88.9) 9/9 (100) 9/9 (100)
0/9 (0) NDb 6/9 (66.7) 3/9 (33.3) ND
0/9 (0) 8/8c (100) 6/9 (66.7) 8/8c (100) 8/9 (88.9)
a
Tortoises (n 5 9) were infected intranasally with 108 CCU of M. agassizii. ND, not done. c Denotes one missing sample. b
were different at all time points p.i. (P , 0.001) (Fig. 1A). Nasal discharge was statistically greater in infected tortoises than in control tortoises at 4 (P 5 0.004), 8 (P 5 0.01), 12 (P 5 0.004), and 16 (P 5 0.01) weeks p.i. (Fig. 1B). Ocular discharge was statistically greater in infected tortoises than in control tortoises at 4 (P 5 0.01), 12 (P 5 0.004), and 16 (P 5 0.005) weeks p.i. (Fig. 1B). Palpebral edema was statistically greater in infected tortoises than in control tortoises at 4 (P 5 0.04), 8 (P 5 0.004), and 12 (P 5 0.04) weeks p.i. (Fig. 1B). Conjunctivitis was statistically greater in infected tortoises than in control tortoises at 12 (P 5 0.04) weeks p.i. (Fig. 1B). The overall cumulative severity of clinical signs increased and then reached a relative plateau (Fig. 1A). However, the individual clinical signs comprising the cumulative scores showed significantly more variability (Fig. 1B). The severity of palpebral edema and conjunctivitis remained relatively constant throughout the 16-week observation period. The severity of nasal and, to a lesser extent, ocular discharge did increase with time following infection (Fig. 1B). Considerable variability in the expression of clinical signs among individual animals occurred (data not shown). Some animals showed a classical plateau response, while others clearly demonstrated intermittent clinical signs. Some individual animals had very severe clinical signs, while others (n 5 3) had clinical signs which had relatively low scores and one animal showed no clinical signs. Infection with M. agassizii resulted in detectable antibody responses by week 8 p.i. (Fig. 2). All of the experimentally infected tortoises seroconverted. Levels of antibody were statistically higher in infected animals than in control animals for all time points .4 weeks p.i. (P , 0.0001) (Fig. 2). No antibody response was detected in any control animal at any time point. Clinical disease outcome: dose-response study. The numbers of M. agassizii used to infect the tortoises initially did not influence the clinical expression of URTD. In most instances, the most severe clinical signs were observed at 8 weeks p.i., regardless of the infective dose. As was seen in the earlier infection trial, there was considerable variability in the expression of clinical signs by individual animals (data not shown). The antibody response (Fig. 3), like the expression of clinical signs, was not affected by the infection dose used. The antibody response in infected animals was first detectable at 6 weeks p.i. and was statistically different from that in the control animals at all time points thereafter (P , 0.001). Histology: experimentally infected tortoises. The nasal cavities of control tortoises consisted of a dorsal multilayered olfactory epithelium (Fig. 4A) and a ventral mucous epithelium consisting of mucus cells intercalated with ciliated epithelial cells (Fig. 4B). Eight of nine experimentally infected tortoises had changes in the nasal epithelia and submucosa (Fig.
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FIG. 2. Specific antibody to M. agassizii in gopher tortoises experimentally infected with M. agassizii (■) and control gopher tortoises (h). Levels of antibody were higher in infected animals than in control animals for all times .4 weeks p.i. (P , 0.0001).
5). The epithelium was intact in all nine tortoises, and no ulcerations were present. One tortoise had mild to moderate changes, five tortoises had moderate changes, and three tortoises had changes that were characterized as moderate to severe. Histology: dose-response study. Changes were observed in the nasal cavity epithelia of experimentally infected tortoises. Eight tortoises were selected for full necropsy. The changes
FIG. 1. Clinical signs of URTD in gopher tortoises experimentally infected with M. agassizii. Tortoises (n 5 9) were infected intranasally with 108 CCU of M. agassizii. No clinical signs were seen for control tortoises (n 5 10), which received sterile broth. (A) Results are expressed as the mean cumulative score, which was calculated as the nasal discharge score plus the mean for the three ocular scores. Infected tortoises (F) had greater cumulative signs than control tortoises (E) at all time points p.i. (P 5 0.001). (B) Results are expressed as the mean clinical sign score 1 standard error on a scale of from 0 (none) to 3 (severe). At 4 weeks p.i., nasal (P 5 0.004) and ocular (P 5 0.01) discharges as well as palpebral edema (P 5 0.04) were greater in infected tortoises than in control tortoises. At 8 weeks p.i., nasal discharge (P 5 0.01) and palpebral edema (P 5 0.004) were greater in infected tortoises than in control tortoises. At 12 weeks p.i., nasal (P 5 0.004) and ocular (P 5 0.004) discharges as well as palpebral edema (P 5 0.04) and conjunctivitis (P 5 0.04) were greater in infected tortoises than in control tortoises. No clinical signs were seen for control tortoises (n 5 10), which received sterile broth (data not shown). The clusters of four bars for each week represent palpebral edema, conjunctivitis, nasal discharge, and ocular discharge from left to right, respectively.
FIG. 3. Specific antibody to M. agassizii in gopher tortoises experimentally infected with different doses of M. agassizii (1, low dose; o, medium dose; ■, high dose) and control gopher tortoises (h).
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FIG. 4. Photomicrograph of the nasal cavity of a control gopher tortoise. (A) Area of mucous and ciliated epithelial cells (M) overlying a lamina propria submucosa (S) primarily consisting of connective tissue and small vessels. (B) Area of multilayered olfactory mucosa (O) overlying a lamina propria submucosa (S) consisting of connective tissue, vessels, and melanophores. Hematoxylin and eosin stain was used.
observed, like the clinical signs, were variable and appeared to be independent of infective dose. In at least one tortoise, changes were different in the right versus left nasal cavity, suggesting that intra-animal variation also occurred. In the
high-dose group (n 5 2), one animal had mild to moderate inflammatory changes; the other tortoise showed moderate changes. Three tortoises in the medium-dose group were selected for necropsy. One tortoise that received the medium
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FIG. 5. Representative moderate to severe changes observed in the upper respiratory tracts of experimentally infected gopher tortoises. The epithelium (E) is hyperplastic, and there are diffuse accumulations of mixed inflammatory cells (IC) in the lamina propria. Hematoxylin and eosin stain was used.
infective dose had moderate changes, and two tortoises in this group had moderate to severe changes. Three tortoises in the low-dose group were selected for necropsy. One tortoise that received the low infective dose had moderate changes, one tortoise in this group had moderate to severe changes, and the final tortoise in this group had severe changes. DISCUSSION The most stringent requirements for definitive proof of a causative relationship between an infectious agent and a disease is the fulfillment of the Henle-Koch-Evans postulates (10, 11). In a free-ranging, wild animal which is also legally protected, this is a daunting challenge. The M. agassizii isolate used in these studies was obtained from an animal with clinical disease. In the present experimental infection studies, this isolate was cultured in vitro and was inoculated into clinically healthy animals which were free of mycoplasma infection for a period of several months as determined by culture, PCR, and serology. The experimentally infected animals developed clinical signs of disease (eight of nine animals) and produced specific antibody against the infectious agent (nine of nine animals). Both the appearance of clinical disease and the seroconversion occurred within reasonable time periods relative to the time of inoculation of the infectious agent. The lesions
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observed in the animals with the experimental infection were similar to those observed both in desert tortoises with natural and experimental infections (4, 16) and in gopher tortoises with natural infections (unpublished data). M. agassizii was recovered from the experimentally infected animals at various times p.i. Thus, we have fulfilled the Henle-Evans-Koch’s postulates and conclude that M. agassizii causes URTD in the gopher tortoise. In this study, the patterns of the experimental infections as well as of the natural infections of gopher tortoises that were observed are consistent with existing knowledge of mycoplasmal respiratory infections in other species. The variability observed in the clinical expression of disease among individual animals is common with other mycoplasmal respiratory infections (32). In most animals, respiratory mycoplasmosis is typified as a slowly progressing, chronic, and seemingly clinically silent infection which may be exacerbated by environmental factors, stress, or other microbial agents (5, 29, 32, 35). Most hosts have difficulty in eliminating the mycoplasma, even in the presence of a strong immune response. In fact, the host immune response is critical for the development of lesions (32). Although overt clinical signs may be inapparent, lesions can range from microscopic to gross, with eventual loss of the normal respiratory epithelium architecture (4, 5, 16, 32). The increased numbers of inflammatory cells, particularly in foci, and the lymphoid hyperplasia observed in the lesions of experimentally infected gopher tortoises are consistent with respiratory mycoplasmosis in other species, including the desert tortoise (16, 32). The relative insensitivity of PCR versus the sensitivity of culture was somewhat surprising. Electron micrographic studies have identified preferentially colonized sites on the mucosal surfaces of ventrolateral depressions in tortoise upper respiratory passages, which may not be accessible by swabbing or lavage (data not shown). Since a prolonged incubation is required for culture, small numbers of M. agassizii may be expanded to a detectable level as a result of microbial growth. Despite its relative insensitivity, PCR still can play an important role in the diagnosis of URTD. A positive PCR result can be obtained with broth cultures after 24 to 48 h of growth, even though the initial PCR with the lavage specimen was negative. An additional problem encountered in diagnosis is the quality of samples. Since gopher tortoises are burrowing animals and many sample collections are done in the field under less than ideal conditions, the samples are often contaminated with bacteria and fungi. Many of these, especially fungi, rapidly overgrow the cultures or alter the medium beyond the pH range tolerated by mycoplasmas for growth. Culture in SP4 broth for 48 h before taking an aliquot for PCR enhances detection of viable mycoplasma, and contamination with other sources of DNA does not interfere with the PCR (data not shown). Isolation of M. agassizii from broth or agar can take up to 6 weeks. If broth cultures are grown for 24 to 48 h and then tested by PCR, we can detect positive culture samples faster, which may be of primary importance if animals are being quarantined or held prior to relocation as part of recommended health surveillance to prevent the spread of disease. A wide variety of potential virulence factors have been suggested for mycoplasmas, including superantigen production, surface antigenic variation, and host immunomodulation (32). It is not uncommon to see a wide variety in the virulence of different strains of the same species within a specific host (7, 8, 17, 28). The strain selected for our studies was obtained from a very ill, naturally infected tortoise. This particular strain appears to be highly virulent, as evidenced by the fact that initial infective doses of only 10 CFU were capable of causing
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both clinical disease and severe lesions. We have preliminary evidence that other strains of M. agassizii do not cause overt clinical disease, even when relatively high infective doses are used. These observations of strain variability are similar to those observed for respiratory mycoplasmosis in rodents and poultry (7, 8, 28). For the most part, any given mycoplasmal species has a relatively limited range of host specificity. Because M. agassizii has a limited temperature range for growth and does not grow at 35°C (3a), it is highly unlikely that it represents a threat to humans or other mammals. Conversely, evidence suggests that other chelonians (turtles and tortoises) may be susceptible to M. agassizii (15, 16, 19, 31). In the past several years, we have seen clinical cases of URTD in tortoises and turtles from zoological collections as well as private collections. It is not uncommon for these reptiles to be housed together, without quarantine or determination of health status. In at least one instance, confiscated star tortoises were sent to a zoo and were later found to have URTD. We have demonstrated the presence of M. agassizii in these animals by serology, culture, and PCR. Treatment with antibiotics alleviates clinical signs but does not eliminate the infection (14a). Appropriate quarantine, screening, and health surveillance of reptiles in collections will be needed to protect animals from URTD. This is particularly important when confiscated shipments of animals with unknown health histories are distributed to zoological settings. Both the gopher and desert tortoises have, to various extents, legal protection due to their decreasing populations. In many cases, the management tools used are relocation of animals to wildlife sanctuaries or other suitable habitats. Until recently, wildlife management decisions have not included considerations of infectious diseases and the possible impact of these diseases on population health. The full impact of relocation efforts involving ill animals is yet to be determined and will require long-term monitoring studies. Research on the impact of mycoplasmosis on wildlife has been limited, but reports of recent disease outbreaks in different wildlife species are provoking interest in mycoplasmosis as a newly emerging (or at least newly recognized) disease threat for wildlife. The most publicized disease outbreak has been seen in Mycoplasma gallisepticum infection of house finches, goldfinches, and blue jays (21, 23, 27). In 1993 an epizootic of polyarthritis occurred in juvenile farmed crocodiles (Crocodylus niloticus) in Zimbabwe (18, 25). A mycoplasma was isolated from the joints and lungs of affected crocodiles, was determined to be a previously unrecognized species, and was named Mycoplasma crocodyli (18, 25). In 1995, a systemic infection of captive adult American alligators at a private facility in Florida was associated with a different species of mycoplasma that had also been previously unrecognized. Unlike the outbreak in crocodiles, the disease in alligators was characterized by a very high mortality rate (.70%) and widespread dissemination of the infectious agent within the tissues of infected animals (3). Infectious diseases are an ever present risk to wildlife, particularly during situations in which animals are removed from their natural habitat for captive breeding programs or during conditions of stress such as release into new habitats, translocation, ecosystem perturbation, and encroachment on their habitats by urbanization (13, 24, 26, 27). Infectious diseases, their implications for population health, and their impact on the success of conservation and management plans are rarely considered in management issues. URTD is an excellent example of the importance of wildlife diseases in population biology. Coupled with habitat destruction and environmental stress factors such as drought, this disease is believed to be a major factor in declines of desert tortoise populations in the
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Mojave Desert (1, 15, 16). Increasing awareness of the role of infectious diseases has resulted in inclusion of disease monitoring and assessment of population health in management decisions in both the desert and tortoise populations. ACKNOWLEDGMENTS This work was supported by a grant from the Walt Disney World Corporation. We acknowledge the technical assistance of Diane Dukes, Michael Lao, Alyssa Whitmarsh, John Hutchison, and Sylvia Tucker. REFERENCES 1. Berry, K. H. 1997. Demographic consequences of disease in two desert tortoise populations in California, USA, p. 91–99. In J. Van Ebbema (ed.), Proceedings: conservation, restoration, and management of tortoises and turtles—an international conference. Wildlife Conservation Society Turtle Recovery Program and the New York Turtle and Tortoise Society, New York, N.Y. 2. Brown, D. R., B. C. Crenshaw, G. S. McLaughlin, I. M. Schumacher, C. E. McKenna, P. A. Klein, E. R. Jacobson, and M. B. Brown. 1995. Taxonomy of the tortoise mycoplasmas Mycoplasma agassizii and Mycoplasma testudinis by 16S rRNA gene sequence comparisons. Int. J. Syst. Bacteriol. 45:348–350. 3. Brown, D. R., M. B. Brown, and J. G. Tully. Comparison of mycoplasma isolated from alligators and crocodiles, abstr. G-10, p. 281. In Abstracts of the 97th General Meeting of the American Society for Microbiology 1997. American Society for Microbiology, Washington, D.C. 3a.Brown, M. B. Unpublished data. 4. Brown, M. B., I. M. Schumacher, P. A. Klein, K. Harris, T. Correll, and E. R. Jacobson. 1994. Mycoplasma agassizii causes upper respiratory tract disease in the desert tortoise. Infect. Immun. 62:4580–4586. 5. Cassell, G. H., W. A. Clyde, and J. K. Davis. 1985. Mycoplasmal respiratory mycoplasmosis, p. 69–107. In S. Razin and M. F. Barile (ed.), The Mycoplasmas, vol. IV. Academic Press, Inc., New York, N.Y. 6. Cox, J., D. Inkley, and R. Kautz. 1987. Ecology and habitat protection needs of gopher tortoise (Gopherus polyphemus) populations found on lands slated for large-scale development in Florida. Technical report no. 4. Florida Game and Fresh Water Fish Commission Nongame Wildlife Program, Tallahassee. 7. Davidson, M. K., J. K. Davis, J. R. Lindsey, and G. H. Cassell. 1988. Clearance of different strains of Mycoplasma pulmonis from the respiratory tract of C3H/HeN mice. Infect. Immun. 56:2163–2168. 8. Davidson, M. K., J. R. Lindsey, R. F. Parker, J. G. Tully, and G. H. Cassell. 1988. Difference in virulence for mice among strains of Mycoplasma pulmonis. Infect. Immun. 56:2156–2162. 9. Diemer, J. E. 1986. The ecology and management of the gopher tortoise in the United States. Herpetologica 42:125–133. 10. Evans, A. S. 1976. Causation and disease: the Henle-Koch postulates revisited. Yale J. Biol. Med. 49:175–195. 11. Evans, A. S. 1977. Limitations of Koch’s postulates. Lancet ii:1277–1278. 12. Hansen, K. L. 1963. The burrow of the gopher tortoise. J. Fl. Acad. Sci. 26:353–360. 13. Hutchins, M., and U. S. Seal. 1991. The role of veterinary medicine in endangered species conservation. J. Zoo Wildl. Med. 22:277–281. 14. Jackson, D. R., and E. G. Milstrey. 1989. The fauna of gopher tortoise burrows, p. 86–98. In J. E. Diemer, D. R. Jackson, J. L. Landers, J. N. Layne, and D. A. Wood (ed.), Gopher Tortoise Relocation Symposium proceedings. Technical report no. 5. Florida Game and Fresh Water Fish Commission, Tallahassee. 14a.Jacobson, E. R. Unpublished data. 15. Jacobson, E. R., M. B. Brown, I. M. Schumacher, B. R. Collins, R. K. Harris, and P. A. Klein. 1995. Mycoplasmosis and the desert tortoise (Gopherus agassizii) in Las Vegas Valley, Nevada. Chel. Cons. Biol. 1:280–284. 16. Jacobson, E. R., J. M. Gaskin, M. B. Brown, R. K. Harris, C. H. Gardiner, J. L. LaPointe, H. P. Adams, and C. Reggiardo. 1991. Chronic upper respiratory tract disease of free-ranging desert tortoises, Xerobates agassizii. J. Wildl. Dis. 27:296–316. 17. Jones, G. E., J. S. Gilmour, and A. G. Rae. 1982. The effects of different strains of Mycoplasma ovipneumoniae on specific pathogen-free and conventionally-reared lambs. J. Comp. Pathol. 92:267–272. 18. Kirchhoff, H., K. Mohan, R. Schmidt, M. Runge, D. R. Brown, M. B. Brown, C. M. Foggin, P. Muvavirirwa, H. Lehmann, and J. Fossdorf. Mycoplasma crocodyli sp. nov., a new species from crocodiles. Int. J. Syst. Bacteriol. 47:742–746. 19. Lawrence, K., and J. R. Needham. 1985. Rhinitis in long-term captive Mediterranean tortoises (Testudo graeca and T. hermanni). Vet. Rec. 117:662– 664. 20. Levell, J. P. 1995. A field guide to reptiles and the law. Serpent’s Tale, Excelsior, Minn. 21. Ley, D. H., J. E. Berkhoff, and J. M. McLaren. 1996. Mycoplasma gallisepticum isolated from house finches (Carpodacus mexicanus) with conjunctivitis. Avian Dis. 40:480–483.
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