INTRODUCTION

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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 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

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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 alcohol-dampened 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 = 10) were sham inoculated intranasally with 0.5 ml (0.25 ml/nostril) of sterile SP4 broth. Tortoises in the experimental infection group (n = 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 80°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 = 6), medium (10³ CCU; n = 6), or high (10⁵ CCU; n = 5) dose of M. agassizii. Tortoises in the control group (n = 10) were sham inoculated intranasally with 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-µl aliquot of the lavage sample was used for PCR analysis; the remaining sample was serially diluted 10-fold to 10² 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 × g for 60 min at 4°C, and the supernatant was aspirated. The pellets were resuspended in 3 to 4 µl of 20 mg of proteinase K (Sigma, St. Louis, Mo.) per ml in 20 µl 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 µl of each sample was removed and was added to 45 µl of a reaction solution containing two primers for the 16S rRNA gene at 1 µM each, deoxynucleoside triphosphates at 200 µM, 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 [5'-CCTATATTATGACGGTACTG-3']) and a Mycoplasma genus-specific region (anti-sense strand nt 1055 to 1031 [5'-TGCACCATCTGTCACTCTGTTAACCTC-3']) (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 µl of product with 2 µl 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 X174 DNA, was included on each gel.

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 µl of a whole-cell lysate of M. agassizii 723 at 10 µg/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 µl of PBST containing 5% nonfat dry milk (PBS-TM) per well. Following washing, 50 µl 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 µl (per well) of a biotinylated monoclonal antibody (monoclonal antibody HL673) against the light chain of desert tortoise immunoglobulins Y and M at 1 µg/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, Calif.) at 1:2,000 in PBS-AZ was added at 50 µl/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 µl/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.

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 (two-group 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

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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.

View larger version (20K): [in this window] [in a new window]   FIG. 1.   Clinical signs of URTD in gopher tortoises experimentally infected with M. agassizii. Tortoises (n = 9) were infected intranasally with 108 CCU of M. agassizii. No clinical signs were seen for control tortoises (n = 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 () had greater cumulative signs than control tortoises () at all time points p.i. (P = 0.001). (B) Results are expressed as the mean clinical sign score + standard error on a scale of from 0 (none) to 3 (severe). At 4 weeks p.i., nasal (P = 0.004) and ocular (P = 0.01) discharges as well as palpebral edema (P = 0.04) were greater in infected tortoises than in control tortoises. At 8 weeks p.i., nasal discharge (P = 0.01) and palpebral edema (P = 0.004) were greater in infected tortoises than in control tortoises. At 12 weeks p.i., nasal (P = 0.004) and ocular (P = 0.004) discharges as well as palpebral edema (P = 0.04) and conjunctivitis (P = 0.04) were greater in infected tortoises than in control tortoises. No clinical signs were seen for control tortoises (n = 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.

View larger version (15K): [in this window] [in a new window]   FIG. 2.   Specific antibody to M. agassizii in gopher tortoises experimentally infected with M. agassizii () and control gopher tortoises (). Levels of antibody were higher in infected animals than in control animals for all times >4 weeks p.i. (P < 0.0001).

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).

View larger version (29K): [in this window] [in a new window]   FIG. 3.   Specific antibody to M. agassizii in gopher tortoises experimentally infected with different doses of M. agassizii (, low dose; , medium dose; , high dose) and control gopher tortoises ().

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. 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.

View larger version (137K): [in this window] [in a new window]   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.

View larger version (141K): [in this window] [in a new window]   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.

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 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 = 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 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

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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 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 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 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.