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Upper respiratory tract disease in gopher tortoises, Gopherus polyphemus

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Upper respiratory tract disease in gopher tortoises, Gopherus polyphemus pathology, immune responses, transmission, and implications for conservation and management
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Mclaughlin, Grace Sheryl
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ix, 110 leaves : ill. ; 29 cm.

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Antibodies ( jstor )
Diseases ( jstor )
Eggs ( jstor )
Female animals ( jstor )
Gophers ( jstor )
Infections ( jstor )
Lesions ( jstor )
Mycoplasma ( jstor )
Tortoises ( jstor )
Wildlife ( jstor )
Dissertations, Academic -- Wildlife Ecology and Conservation -- UF ( lcsh )
Gopher tortoise -- Diseases ( lcsh )
Mycoplasma diseases in animals ( lcsh )
Wildlife Ecology and Conservation thesis, Ph. D ( lcsh )
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bibliography ( marcgt )
theses ( marcgt )
non-fiction ( marcgt )

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Thesis (Ph. D.)--University of Florida, 1997.
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Includes bibliographical references (leaves 97-109).
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Also available online.
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Typescript.
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Vita.
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by Grace Sheryl McLaughlin.

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UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES,
GOPHERUS POLYPHEMUS:
PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION,
AND IMPLICATIONS FOR CONSERVATION AND MANAGEMENT








BY

GRACE SHERYL MCLAUGHLIN



















A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

1997



















DEDICATION


This dissertation, the years that went into it, and the drive for knowledge
that underpinned it, are a result of and dedicated to, my parents:


Grace Slater Gibson "Billie" McLaughlin 1928-1978


and


Robert Douglas McLaughlin 1925-1994




Thank you.









ACKNOWLEDGMENTS



I thank Dr. Don Forrester for accepting me as a graduate student, and for his support and guidance through the years. Drs. Kathy Ewel and Paul Gibbs assisted my growth scientifically and professionally in the first half of my program. Dr. Ewel's support was instrumental in obtaining my fellowship, and Dr. Gibbs was responsible for a trip to Australia. Drs. Mel Sunquist and Wiley Kitchens weathered the changes in my project with grace and humor, and Dr. Kitchens was especially helpful in teaching me to argue my positions and not back down when I knew I was right. It took several years for Dr. Mary Brown to get me into her lab, and her support in presenting me to her colleagues is appreciated. Dr. Elliott Jacobson has done his best to teach me clinical pathology and histopathology and has been very supportive of my contributions to the overall project. Dr. Paul Klein has given me some valuable insights into the critical thinking process.

Without my co-workers Drs. Dan Brown and Isabella Schumacher, Sylvia Tucker, Barbara Crenshaw, and Cathie McKenna, and technicians Alyssa Whitemarsh, Michael Lao, and Dave Bunger, this research would have been impossible. I benefited from Dan's, Isa's and Barb's teaching abilities, and their willingness to discuss theory, practical applications, philosophical underpinnings and differences of opinion. Mr. Clement Lindsey and his staff cared for my research animals.

I thank Dr. Tim Gross for all his help with eggs and hatchlings, and John Wiebe and Carla Weiser for their care. Drs. Dale Jackson and Michael Ewert provided advice. Drs. Bruce Homer and Claus Buergelt have given me more strength and support than they




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knew and taught me much. Many of the VMTH personnel, especially Neil, Danielle, Tom, Dana, and An have assisted and taught me, and cheered me up when I needed it.

Ms. Joan Berish has provided an incredible amount of advice, support, and

friendship throughout the eight years I have been studying gopher tortoises, and I only hope to reciprocate. Garry Foster's assistance through my early years at UF were invaluable, as were the support of my fellow graduate students in Dr. Forrester's lab, Marisol Sepulveda and Don Coyner. My brother Mark, my "sister" Megan, and friends Vicky, Bill, Sharon, Kay, Pierre, Andrea, and Tania helped me get here and stick it out, and deserve thanks for their support. There have been several other people who have helped me at various times and in different ways, especially those of the gopher tortoise sodality, and my thanks go to them also. I also thank the members and friends of the Unitarian Universalist Fellowship of Gainesville for being my family and caring for me.

I was supported by a Presidential Research Fellowship and a Gatorade Grant from the University of Florida, and funds from The Walt Disney World Company.





















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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ......................................... iii

ABSTRACT ................................................... viii

CHAPTERS

1 INTRODUCTION ....................................... 1

Gopher Tortoise Natural History ............................ 1
Tortoises and Upper Respiratory Tract Disease ................... 6
Mycoplasmal Respiratory Diseases in
Domestic Animals and Humans .......................... 10
Mycoplasmal Diseases of Wildlife ................ ............ 13
Chronic Manifestations of Mycoplasmal Infections ............... 18
Project Overview and Specific Objectives ...................... 18

2 METHODS............................................ 21

Tortoises, Intake Procedures, Clinical Assessments
and Sampling Methods ................................ 21
Culture Procedures ....................................... 22
PCR Procedure ......................................... 22
ELISA Procedure ........................................ 24
Study Group Assignment .................................. 25
Husbandry Procedures ................................... 26
Necropsy Procedures ................................... 27
Histopathology Procedures ................................. 28
Statistical Analyses ...................................... . 29

3 NATURALLY OCCURRING UPPER
RESPIRATORY TRACT DISEASE ...................... 30

Methods ........................................ 30
Tortoises ........................................... 30
Necropsy and Histology Procedures ................ ....... 31
Microbial Isolation .................................... 31
Electron Microscopy .......... ........................ 32
Results ................................................ 33
Normal Anatomy and Histology ......................... 33
Pathologic Findings ................................... 37
ELISA and PCR Results ................................ 40
Microbial Isolation Results .............................. 40
Discussion ............................................ 40

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4 EFFECTS OF REPEATED EXPOSURE ON
SEROPOSITIVE ADULTS ............................ 46

Introduction ............................................. 46
Methods ........................................ 47
Statistical Analyses ...................... ............. 47
Experimental Design ..................................... 47
Results ........................................ 49
Clinical Signs ........................................ 49
Culture and PCR Results ................................ 53
ELISA Results .................. .................... 53
Histology Results .................................... 55
Discussion ................................................ 55

5 HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII .. 59

Introduction ............................................... 59
Methods ........................................ 59
Experimental Design ................................... 60
Results ........................................ 64
General Observations .................................. 64
Evidence of Transmission ofMycoplasma agassizii ............ 66
Transmission Probabilities ............................... 68
Discussion ............................................. 71

6 VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII ..... 75

Introduction .......................................... 75
Methods .............................................. 75
Egg Collection and Incubation ........................... 76
Culture and PCR Procedures ............................ 76
ELISA Procedures .................................... 77
Results ........................................ 78
Clutch Sizes, Fertility and Hatching Rates ................... 78
Culture and PCR Results ................................ 79
ELISA Results ....................................... 80
Discussion ........................................... 82

7 ENVIRONMENTAL TRANSMISSION OF
MYCOPLASMA AGASSIZII .......................... 84

Introduction ............................................... 84
Methods ........................................ 84
Results .............................................. 85
D iscussion ............................................. 86



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8 CONSERVATION AND MANAGEMENT IMPLICATIONS
OF UPPER RESPIRATORY TRACT DISEASE ............. 88

Implications for Conservation and Management ................. 88
Establishing Goals .................................... 89
Understanding URTD and Test Results ..................... 89
Developing Questions and Conducting Surveys
or Monitoring Programs ............................. 91
Weighing Management Options and
Formulating Management Plans ........................ 93
Summary of Conservation and Management Implications ....... 95
Further Research ....................................... 96


LITERATURE CITED ............................................. 97

BIOGRAPHICAL SKETCH ........................................ 110




































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Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES, GOPHERUS POLYPHEMUS: PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION, AND IMPLICATIONS FOR CONSERVATION AND MANAGEMENT By

GRACE SHERYL MCLAUGHLIN

May 1997

Chair: Donald J. Forrester, Ph. D.
Cochair: Mary B. Brown, Ph. D.
Major Department: Wildlife Ecology and Conservation


Upper respiratory tract disease (URTD) of tortoises is caused by the mollicute Mycoplasma agassizii, and is characterised by nasal and ocular discharge, palpebral edema, and conjunctivitis. Hyperplasia and dysplasia of the nasal passage and cavity epithelia and inflammatory infiltrates are seen histologically. In order to provide data for management decisions and to better understand URTD, I studied uninfected, naturally infected, and experimentally infected tortoises. The pathological and immune responses of tortoises to and transmission of M agassizii were investigated using clinical and histological observations, culture and polymerase chain reaction (PCR) tests, and an enzyme-linked immunosorbent assay (ELISA). Infection with M. agassizii caused mild to severe damage to mucosal and olfactory nasal epithelia, with increased damage in longerviii










term infections. Although clinically healthy, ELISA-positive, culture and PCR-negative tortoises may have eliminated the bacteria, when five such animals were examined at necropsy, three were found to harbor M. agassizii in the nasal cavities. When seropositive tortoises were challenged with M. agassizii, a more rapid and more severe clinical response resulted than on initial exposure, and plasma antibody levels began rising more quickly. When uninfected animals were housed with infected individuals, horizontal transmission occurred, probably via direct contact, but possibly via food, water, or fomites. Transmission was more likely to occur from a tortoise that was clinically ill and culture or PCR-positive. There was no discernible transmission when tortoises inhabited pens or entered burrows previously occupied by ill tortoises. There was no demonstrable vertical transmission, although there was transfer of maternal antibodies via egg yolk. The level of antibodies in egg yolk or hatchling plasma was approximately 10-20% of that in maternal plasma. Movement of tortoises during relocation, repatriation, or restocking efforts potentially could transport M. agassizii to previously uninfected sites. Because of the uncertainty involved in determining latent infections, ELISA-positive animals should not be moved to locations with no seropositive individuals. However, they can be used in captive breeding efforts and their offspring released into the wild. Clinically ill animals should not be relocated to new sites, and should not be housed with clinically healthy animals in temporary holding situations, such as on-site relocations, or in captive breeding programs.








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CHAPTER 1
INTRODUCTION



Gopher Tortoise Natural History


Gopher tortoises, Gopherus polyphemus, are found in the southeastern United States, on the coastal plain from southern South Carolina south through Georgia and throughout Florida, and west through southern Alabama, Mississippi, and Louisiana. The major population concentrations are in Florida and southern Alabama and Georgia, with only remnant populations in South Carolina, Mississippi and Louisiana (Auffenberg and Franz 1982, Diemer 1992a). Populations are concentrated in areas with deep sandy soils suitable for digging. Vegetation associations in which tortoises are found include longleaf pine-xerophytic oak woodlands, palmetto scrub, sand pine scrub, oak scrub, beach scrub, coastal strands, pine flatwoods, dry prairies, native pasture, and savanna, as well as ruderal habitats (Landers and Speake 1980, Lohoefener and Lohmeier 1981, McRae et at 1981, Campbell and Christman 1982, Diemer 1986, Breininger et al 1988).

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 (Eisenberg 1983). Gopher tortoises live in loose colonies, with considerable movement of tortoises among groups over the years (Diemer 1992b). Colonies may be defined more by the availability of suitable soils for digging burrows, or the distribution of food resources, than by social



1







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interactions (Campbell and Christman 1982). Gophers are the most fossorial of the four North American species of tortoises, digging burrows that may extend 5 meters down from the surface and 15 meters in length (Hansen 1963, Diemer 1986). The burrows provide a microclimatically stable environment for not only the tortoises, but also for numerous commensals. Approximately 60 vertebrate species--from snakes to birds--and over 300 invertebrates--including spiders, crickets, and beetles--have been found in tortoise burrows or observed using them as permanent homes or refuges from heat, cold, fire, and predators (Jackson and Milstrey 1989, Lips 1991, Witz et al. 1991). Some invertebrate species are obligate commensals, and occur only in active tortoise burrows, where they feed on tortoise feces and other invertebrates and, in turn, are eaten by other commensals, such as gopher frogs (Rana areolata) and Florida mice (Podomys floridanus) (Woodruff 1982). Several species that exclusively or frequently use tortoise burrows have legal protection in Florida and other parts of their ranges. These include scarab beetles (F. Scarabaeidae), indigo (Drymarchon corals couperi) and pine (Pituophis melanoleucus) snakes, gopher frogs, mole skinks (Eumeces egregius), burrowing owls (Athene cuniculariafloridana), and Florida mice (Cox et al. 1987).

The soil disturbance resulting from the digging of the burrows allows deeper

access of air and water into the soil profile, as well as providing bare mineral soil patches on the surface. When a burrow is abandoned the soil mound, or apron, in front of the entrance no longer undergoes continual disturbance, allowing certain plants to colonize the area. The composition of the plant assemblage on the mound may differ from that in the surrounding undisturbed area, providing a mosaic of small patches in the habitat (Breininger et al 1988, McLaughlin 1990). Tortoises consume a wide variety of grasses






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and forbes, and readily ingest the fruits of many shrubs when available (Garner and Landers 1981, Macdonald and Mushinsky 1988). Subsequent tortoise movements may help spread the seeds to suitable habitat, and some seeds (e.g., gopher apple) may germinate more readily after passage through the tortoise gut.

Gopher tortoises are long-lived, slow to mature, and have a low reproductive rate. Age estimates extend up to 100 yr, although 60-80 yr is considered a more reasonable estimate. Age to sexual maturity ranges from 10 to 20 yr, with possible latitudinal association (Alford 1980, Iverson 1980, Landers et al. 1982, Wright 1982, Doonan 1986, McLaughlin 1990, Mushinsky et al. 1994). Gopher tortoises lay one clutch of eggs annually, generally ranging from 2-10 eggs in size, with the average being 4-8 (Dietlein and Franz 1979, Iverson 1980, Landers et al. 1980, Linley and Mushinsky 1994). In some areas with excellent food resources and large tortoises, clutch sizes may average 9-10, with a range of 6-14 eggs (McLaughlin, 1990). Although some wild individuals have produced over 20 eggs in one clutch (Godley 1989, L. Macdonald, personal communication), this is very unusual. With predation rates of up to 95% on eggs and hatchlings, and further high losses ofjuveniles aged 1-5 yr, less than 10%, and possibly as low as 1%, of eggs laid eventually produce reproductive adults (Douglass and Winegarner 1977, Auffenberg and Iverson 1979, Alford 1980, Iverson 1980, Landers et al. 1982, Diemer 1986, Wilson 1991).

The gopher tortoise is listed in Appendix II of the Convention on International Trade in Endangered Species of Wild Fauna and Flora (C.I.T.E.S.), which requires permits for the exportation of the species from the U. S. to any signatory nation, or for reexportation (Levell 1995). However, if federal or state regulations are more restrictive






4


than C.I.T.E.S., those take precedence. Legal protection is extended to the species in all states within the range, although the levels of protection vary. The populations west of the Tombigbee and Mobile Rivers in Alabama, Mississippi and Louisiana are on the federal threatened species list, which prohibits the taking, exportation, or interstate movement of individuals originating from that region without permit, and the possession, transportation, purchase, and/or sale of illegally obtained specimens. Permits issued by the U. S. Department of Interior can be obtained for scientific research on wild populations, and possession of limited numbers of individuals for exhibition, education, and/or research. State permits are required also. The gopher tortoise is listed in Alabama as a protected nongame species, in Mississippi as endangered, and in Louisiana as threatened. Louisiana regulations also prohibit the use of gasoline, chemicals, or volatile substances to flush reptiles from burrows or other hiding places. Georgia lists the species as threatened and issues scientific collection permits only to qualified institutions and individuals for educational or research purposes. Although the law states that burrows may not be disturbed nor destroyed, nor may explosives, chemicals or smoke be introduced into them to drive out wildlife, the code explicitly exempts poisonous snakes. Because venomous snakes, particularly rattlesnakes, use gopher tortoise burrows, tortoises can be adversely impacted by such activities. South Carolina was the last state to extend legal protection to gopher tortoises (Mann 1990, Levell 1995), and now lists the species as endangered, with permits required for any activities involving tortoises. In Florida, where the gopher tortoise is listed as a species of special concern (Wood 1996), regulations prohibit taking and disturbing of tortoises and their habitats, although exceptions are granted regularly for development, agriculture, and mining operations. In the past, human consumption of






5


tortoises was a major impact on regional distribution, and current poaching activities may extirpate local aggregations (Diemer 1989, Mann 1990). Large scale conversion of longleaf pine habitats to slash pine plantations, transformation of native pasture or savanna to "improved" pasture, citrus groves, or row crops, mining operations such as phosphate, mineral sand, and gravel mines, and urban/suburban development are the main threats to continued gopher tortoise survival today (Diemer 1986, Cox et at 1987, Diemer and Moore 1994). Lack of natural fire regimes due to suppression efforts by humans may alter vegetation mosaics in remaining habitat, rendering them less suitable to continued maintenance of tortoise populations (Mushinsky 1986, Mushinsky and Gibson 1991).

Although tortoise numbers increase with increasing areal extent of available habitat, densities remain constant or decrease (Mushinsky and McCoy 1994). Fragmentation of mainland areas, with ever smaller islands of suitable habitat surrounded by agricultural and urban development, may force tortoises into higher density populations than would occur normally. Vegetational changes resulting from reduced fire incidence in small areas, particularly increased canopy closure and decreased herbaceous vegetation (Mushinsky 1985, 1986), can lead to decreases in reproductive rates and juvenile survival (Auffenberg and Franz 1982, McLaughlin 1990, Mushinsky and McCoy 1994). Increased intraspecific interactions at higher densities (McRae et al. 1981) may lead to elevated physiological stress, potentially affecting immune system function and rendering animals more susceptible to disease. Increased densities and number of interactions multiply the opportunities for transmission of communicable agents, making fragmented populations more likely to sustain high morbidity during epizootics. If mortality is high, lowered reproduction due to habitat degradation may be insufficient for population recovery.






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Along with increased physiological stress placed on tortoise populations from human activities, there may be toxicological or immunological stress from chemicals introduced into the environment. No research has been conducted into the effects of herbicides, fungicides, insecticides, and fertilizers on gopher tortoise health, growth, or reproduction. Investigations of disease in free-ranging populations of tortoises have begun only recently.


Tortoises and Upper Respiratory Tract Disease


Although individual captive and wild gopher tortoises have been observed with

clinical signs of respiratory diseases for over 20 yr (E. R. Jacobson, unpublished data), the first documentation of a larger-scale disease outbreak was in 1989, when an epizootic of upper respiratory tract disease (URTD) was documented on Sanibel Island, Lee County, Florida (G. S. McLaughlin and M. Elie, unpub. data). With the loss of 25-50% of breeding-age adults in one population, recovery could take 50-150 yr (G. S. McLaughlin unpub. data), barring further major losses and without substantial habitat improvement leading to increased recruitment.

In the 1980s, large-scale population reductions (33-76% over 10 yr) of desert tortoises (Gopherus agassizii) were documented at several sites in the western Mojave Desert of California and at one site in the eastern Mojave (Corn 1994, Berry in press). Tortoises with clinical signs of URTD were observed among the remaining populations at several sites (Knowles 1989; Berry 1990, in press). As a result of the declines, tortoises in the Mojave Desert north and west of the Colorado River were declared threatened (U.S. Fish and Wildlife Service 1990).






7


Clinical signs of URTD in gopher and desert tortoises include serous, mucoid, or purulent discharge from the nares, excessive tearing to purulent ocular discharge, conjunctivitis, and edema of the eyelids and ocular glands (Jacobson et al. 1991, G. S. McLaughlin personal observations). Individual infected tortoises vary in the suite of signs they have, and the severity can vary from day to day. Nares may become occluded with caseous exudate, preventing externally visible nasal discharge. Lymphocytic infiltration of the corneas, while rare, may decrease an animal's ability to forage or avoid predators. Tortoises may become lethargic and anorectic, leading to dehydration, emaciation, and eventual death from cachexia. Lethargy, nonresponsiveness to stimuli, and altered behavior patterns--such as basking at lower temperatures than normal--may render a tortoise more susceptible to predation. Moribund animals often develop petechial to ecchymotic hemorrhages under the scutes, especially visible on the plastron (G. S. McLaughlin pers. obs.), which may be due to septicemia caused by secondary infection with opportunistic bacteria.

Several agents were hypothesized to cause respiratory tract disease in tortoises, including viruses, Mycoplasma sp. (Lawrence and Needham 1985), and Pasteurella testudinis (Snipes and Biberstein 1982). However, experimental infections to determine the etiologic agent were not conducted. Beginning in 1989, efforts were undertaken to determine the etiology of URTD. Clinically ill and healthy desert tortoises from California were examined and samples collected, either from live animals or at necropsy. Hematologic and serum biochemical evaluations, liver vitamin and metal determinations, and pathologic and microbial investigations were conducted (Jacobson et al. 1991).






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Hemoglobin concentration and phosphorous levels were lower in ill than in healthy tortoises, and ill tortoises had higher sodium, blood urea nitrogen (BUN), creatinine, serum glutamic oxalacetic transaminase (SGOT) activity, and total cholesterol levels. Levels of liver and serum vitamins A and E did not differ between healthy and ill tortoises, nor did liver values of selenium, copper, lead, or cadmium. However, iron and mercury were higher in ill than in healthy tortoises. Healthy tortoises did not exhibit any histological abnormalities in nasal mucosa or other tissues. Ill tortoises had less fat than controls, atrophied thymuses, increased numbers of lymphocytes in the sinusoids of the spleens, and increased amounts of iron granules in the hepatocytes. The most striking and consistent changes were in the nasal passage and cavity tissues (Jacobson et al. 1991). Grossly, passages and cavities of ill tortoises contained moderate to large amounts of serous to purulent exudate. Histologically, the tissues exhibited loss of mucosal glands and goblet cells, proliferation of epithelial cells, infiltrates of lymphocytes and histiocytes, loss of cilia, dysplasia of the olfactory epithelia with infiltration of heterophils, basal cell hyperplasia, occasional squamous metaplasia, and occasional erosion and ulceration of mucosal epithelia. By electron microscopy, pleomorphic organisms resembling Mycoplasma sp. were seen on cell surfaces and tightly adhered to cell membranes of ill tortoises, but not healthy ones. Significantly more Pasteurella testudinis were isolated from ill than healthy tortoises, and Bacillus sp. and Mycoplasma-like organisms were isolated only from ill tortoises. By electron microscopy, the latter resembled those seen on the nasal cavity tissues of ill tortoises (Jacobson et al. 1991).

The mycoplasma was determined to be a new species, provisionally named

Mycoplasma agassizii (Brown et al. 1994). An enzyme-linked immunosorbent assay






9


(ELISA) to detect antibodies against the mycoplasma in plasma and serum samples was developed (Schumacher et al. 1993), and experiments were undertaken to fulfill Koch's postulates. The disease was induced by inoculation of tortoises with pure cultures of the mycoplasma, but not Pasteurella testudinis (Brown et al. 1994). Histologically, the lesions were consistent with those seen in the previously examined naturally infected tortoises. Additional work led to the development of a polymerase chain reaction (PCR) test to detect the bacteria in nasal flush and swab samples (Brown et al. 1995).

Histologic examination of nasal tissues and microbiologic evaluation of flush and swab samples from clinically healthy and clinically ill tortoises from Las Vegas Valley, Nevada, resulted in findings in the ill tortoises similar to those in the California tortoises, with 92% showing lesions of URTD, 50% being culture positive for M. agassizii, and 100% reacting positively in the ELISA (Jacobson et al. 1995). However, 73% of the "healthy" tortoises had lesions consistent with URTD, 50% were culture positive for M. agassizii, and 42% were seropositive for antibodies against the bacteria. These findings demonstrate that the disease can exist in a subclinical form in a substantial proportion of a population. An annual cycle of convalescence and recrudescence of clinical signs has been seen in captive desert (I. M. Schumacher, pers. comm.) and gopher (D. L. Morris, pers. comm.) tortoises. Other mycoplasmal diseases also can exist as chronic, subclinical infections, with recurrence of clinical signs and increases in transmission potential when the host is stressed (see below). Annual fluctuations in temperature, rainfall, and forage availability may be sufficient to cause detectable outbreaks in an infected population. Increased morbidity and mortality may occur in times of unusually severe environmental stress, such as prolonged drought, hurricanes, excessive rainfall with flooding of burrows,






10


or very cold winters. Human impacts on tortoises and their habitat, whether through disruption of normal behavior patterns, degradation of habitat through agriculture, silviculture, mining or development operations, or pollution, may cause sufficient physiological stress to trigger proliferation of the mycoplasma and recurrence of signs. Capturing and transporting of tortoises during relocation, restocking and repatriation efforts also may be significant sources of stress.


Mycoplasmal Respiratory Diseases in Domestic Animals and Humans


Mycoplasmas cause respiratory disease in other taxa, including rodents, poultry, swine, ruminants, and humans. All have similar pathological effects, which are described below, and many are exacerbated by concurrent infections or environmental stress.

Murine respiratory disease caused by Mycoplasma pulmonis has caused problems in laboratory settings for more than 70 yr (Lindsey et al. 1971). It has necessitated complex and expensive control measures, including various types of isolation mechanisms and cesarean delivery of mice and rat pups to reduce the prevalence of M. pulmonis infections in colonies (Cassell et at 1984, Davis et al. 1987). Interactions among environment, host, and strain factors influence the impacts at individual and population levels (Simecka et al. 1992). Histologically, lesions are characterized by lymphoid hyperplasia and chronic inflammation (Cassell et al. 1985).

Poultry respiratory diseases can be caused by viruses, mycoplasmas, and other

bacteria, singly or in combination. Without concurrent viral or other bacterial infections, disease can be mild or subclinical (Kerr and Olson 1967). Environmental factors, particularly dust and ammonia levels, as well as strain differences, affect the severity of









outbreaks of mycoplasmosis. Other stress factors, such as crowding and cold weather, also influence morbidity and mortality (Jordan 1972). Mycoplasma gallisepticum causes chronic respiratory disease in chickens and sinusitis in turkeys, and affects ring-necked pheasants (Phasianus colchicus), chukar (Alectoris chukar), northern bobwhite (Colinus virginianus), common peafowl (Pavo cristatus) (Yoder 1991) and Japanese quail (Coturnixjaponica) (do Nascimento and do Nascimento 1986). Lower respiratory tract complications are seen rarely in turkeys (Simecka et al. 1992). Histologically, hypertrophy and hyperplasia of respiratory epithelia, including mucous cells, are seen, as is lymphoid infiltration of the submucosa. Heterophilic exudate is seen in the airways (Nunoya et al. 1987, Trampel and Fletcher 1981). Mycoplasma meleagridis infection is seen primarily in chicks and poults up to 10 wks of age, and is sexually transmitted. Turkeys, Japanese quail, and peafowl develop air sacculitis, sometimes with accompanying tracheitis and pneumonia, but not sinusitis. Histologically, the lesions are characterized by hypertrophy and hyperplasia of the air sac epithelia, with edema and lymphoid infiltration (Stipkovits 1979). Young chickens and turkeys also are susceptible to M. synoviae, usually in conjunction with Newcastle disease virus or infectious bronchitis (Hopkins and Yoder 1984, Springer et al. 1974). As with M. gallisepticum, turkeys develop sinusitis (Stipkovits 1979). All three bacteria can be transmitted vertically (ie., via the egg) (Simecka et at 1992).

Swine develop mild pneumonia when infected by M hyopneumoniae, and although mortality is virtually nil, the disease is chronic, causing slow growth and reduced weight gains, decreasing profitability (Oboegbulem 1981, Jericho 1986). Although signs can disappear, recurrences will occur with weather changes, viral infections, and other







12


stressors (Whittlestone 1976). Lesions are alveolar, initially characterized by neutrophilic infiltrates and later by lymphocytes and macrophages (Baskerville 1972).

Contagious bovine pleuropneumonia (CBPP), although not present in the United States, is an economically important disease of cattle and water buffalo caused by M mycoides subsp. mycoides (Howard and Gourlay 1978, Trichard et al. 1989). Morbidity rates can reach 100% with up to 50% mortality, and recovered animals maintain a carrier state. Neutrophilic exudate in the airways, serous exudate in the pleural cavity, serofibrinous exudate in alveoli, and edema and necrosis of regional lymph nodes and interlobular septa are seen at necropsy. Interseptal, peribronchial, and perivascular lymphoid infiltration can be seen histologically (Hudson 1971). Calf pneumonia, with high mortality among dairy calves, is caused primarily by viruses, but can be caused by either M. bovis or M. dispar (Stalheim 1983, Bryson 1985). Infection with the former is characterized by peribronchiolar and alveolar monocyte infiltration, while the latter causes interstitial pneumonia with monocyte infiltration of alveolar walls, but not peribronchiolar infiltration. Mycoplasma dispar also causes superficial and asymptomatic infection of respiratory mucosa (Woldehiwet et al. 1990).

Lambs are susceptible to infection with M ovipneumoniae, which is transmitted to neonates from the ewes. Pneumonia, characterized by coughing, sneezing, nasal discharge, fatigue, and poor weight gain, can develop as colostral antibodies wane. Lesions are characterized by alveolar proliferation, nodular lymphoid hyperplasia, and penibronchial and perivascular lymphoid infiltration (Carmichael et al. 1972, Foggie et al. 1976). Severity can be exasperated by infection with Pasteurella haemolytica biotype A (Jones et al. 1982).






13


Goats can suffer 60-100% mortality when infected by M. mycoides subsp.

mycoides, M. mycoides capri, or Mycoplasma strain F38, making contagious caprine pleuropneumonia the most economically important goat disease. Pathologically, signs and lesions are similar to those seen in CBPP, with the addition ofpolyarthritis (Cottew 1984).

Humans are susceptible to infection with M. pneumoniae, which causes

tracheobronchitis and, less commonly, primary atypical, or walking, pneumonia (Chanock et al. 1963, Clyde 1983). The upper respiratory tract is affected, not the alveoli, and otitis media is seen also (Clyde 1979, Mansel et al. 1989). Immunopathological sequelae, although rare, can include arthritis, dermal lesions, cardiopathy, and neurological complications (Johnston et al. 1983, Lyell et al. 1967, Naftalin et al. 1974, Murray et al. 1975). Histologically, perivascular and peribronchial monocytic infiltrations with some neutrophilic exudate are seen (Dajani et al. 1965). Most commonly affected are 5-9 yr old children, with a decline in incidence until about age 25, then an increase in 30-40 yr olds (Denny et al. 1971). The disease is endemic, with slight increases in late summer and fall, and cyclic epidemics occurring at 4-7 yr intervals (Krause and Taylor-Robinson 1992).


Mycoplasmal Diseases of Wildlife


Research on the impact of mycoplasmosis on wildlife is limited, but recent developments are provoking interest. In early 1994, house finches (Carpodacus mexicanus) in the mid-Atlantic and northeastern regions of the United States were found with conjunctivitis, rhinitis, and sinusitis. Many birds were found dead or dying and were submitted to wildlife care facilities, veterinary hospitals, and state agricultural laboratories. Many of those birds had lesions that were histologically compatible with mycoplasma






14


infections, and were infected with M gallisepticum (Ley et al. 1996, Luttrell et al. 1996). Since then, the organism has been associated with morbidity and mortality in other species ofpasserines, including goldfinches (Carduelis tristis) (Nettles 1996), and was transmitted to a blue jay (Cyannocitta cristata) in a rehabilitation facility (Ley et al. 1996). Conjunctivitis and M. gallisepticum also were found in house finches in Georgia (Luttrell et al. 1996). The potential for this organism to spread over a large area due to the long distances traveled by migratory birds, the mixed species flocks in which they congregate, and local concentrations of many species around bird feeders is of great concern to many ornithologists and ecologists, as well as poultry producers. There are indications that the strain, while highly pathogenic to chickens, is not readily transmissible to poultry under natural conditions (Nettles 1996). A new species of mycoplasma (M. sturni) was isolated from the conjunctiva of a European starling (Sturna vulgaris) with conjunctivitis found in Connecticut during the epornitic (Forsyth et al. 1996). Although it was isolated in pure culture, it was described as growing rapidly, whereas pathogenic mycoplasmas typically grow slowly; therefore, it could have overgrown M. gallisepticum if that species had been present. The pathogenicity ofM. sturni needs to be investigated.

Three new species of mycoplasmas have recently been described from raptors in Europe (Poveda et al. 1994). All were associated with respiratory diseases clinically and histologically consistent with those caused by Mycoplasma spp., with lesions including hyperplasia of mucous glands, lymphoid hyperplasia, and perivascular cuffing (Poveda et al. 1990). The species were isolated from buzzards (Buteo buteo), saker falcons (Falco cherrug), and griffon vultures (Gypsfulvus), and have been named, respectively, M buteonis, M. falconis, and M. gypis. Pathogenicity of these species, their distribution in






15

wild, free-ranging birds, and their potential impacts on the populations need to be further investigated.

Several species of mycoplasmas, including M. cloacale (Bradbury et al. 1987,

Goldberg et al. 1995) and M. anatis (Ivanics et al. 1988, Poveda et al. 1990, Goldberg et al. 1995), have been isolated from semi-domestic and wild ducks and other avian species throughout the world. Stipkovits et al. (1986) reported isolation of M. cloacale from geese with inflammation of the cloaca and phallus, but Goldberg et al. (1995) found no association of M. cloacale with disease in wild mallards (Anas platyrhynchos), black ducks (A. rubripes), or canvasbacks (A. valisneria). Stipkovits (1979) reported pathogenicity of M. anatis to domestic ducklings and eggs, and neurological signs have been recorded in ducks infected with M. anatis (Ivanics et al. 1988). Samuel et al. (1995) infected game-farm mallard eggs with M. anatis and found reduced hatchling success, hatchling size and growth rates. Hatchlings infected at 1 d of age did not have slower growth rates. Goldberg et al. (1995) found esophagitis, tracheitis, and vaginitis in female mallards from which M. anatis was isolated, and presented evidence for vertical transmission of M. anatis in a wild gadwall (Anas strepera). Potentially, M. anatis infections could reduce recruitment in wild duck populations. Experimental infection of ducklings with M. gallisepticum resulted in suppressed growth rates (Stipkovits 1979). Four other unidentified mycoplasmas were isolated from mallards, gadwalls, and black ducks, but could not be associated definitively with disease (Goldberg et al. 1995).

Although M. gallisepticum has been isolated from wild turkeys, most cases have occurred in birds with close association to domestic poultry (Davidson et al. 1982, Jessup et al. 1983, Luttrell et al. 1991, Fritz et al. 1992). Even though experimental infection of






16


pen-reared wild turkeys has resulted in decreased productivity (Rocke et al. 1988), there is little evidence to indicate that infection with M. gallisepticum strains commonly occurring in domestic fowl poses a threat to wild turkey populations (Luttrell et al. 1991), or that wild turkey populations are important in the epizootiology ofM. gallisepticum (Davidson et al 1988). Mycoplasma gallopavonis has been isolated from wild turkeys in Texas (Rocke and Yuill 1987), South Carolina, Georgia (Luttrell et al. 1991), Colorado, New Mexico, and Oklahoma (Fritz et al. 1992). Although lethal to experimentally infected domestic turkey eggs (Rocke and Yuill 1987), the pathogenicity ofM. gallopavonis to wild turkeys has not been investigated. Mycoplasma synoviae and other, untyped, Mycoplasma spp. were isolated from turkeys in Arizona (Fritz et al. 1992), but no association with disease was found. As with M. gallisepticum, wild turkeys do not appear to be important in the epizootiology ofM. synoviae or M. meleagridis (Davidson et al. 1988).

In 1993 an epizootic ofpolyarthritis occurred in juvenile farmed crocodiles

(Crocodylus niloticus) in Zimbabwe. The outbreak was characterized by high morbidity, but low mortality. A mycoplasma was isolated, determined to be a previously unrecognized species, and named M. crocodyli (Mohan et. al 1995). In 1995 a die-offof captive adult American alligators (Alligator mississippiensis) at a private facility in Florida, with very high mortality, was associated with systemic infection with a different species of mycoplasma, also previously unrecognized (Brown et al. 1996a). The name M. lacerti has been proposed. In addition to M. agassizii, a second mycoplasma was found in desert tortoises with evidence of URTD, including clinical signs, histologic lesions, and/or positive ELISA tests (Brown et al. 1995). In a small pilot study, that organism was found






17

to cause disease and seroconversion in gopher tortoises previously unexposed to either mycoplasma (D. R. Brown, et al., unpub data).

Mycoplasmas have been isolated from bighorn sheep (Ovis canadensis) with

pneumonia associated with P. haemolytica, and there is concern that these may have a role in the disease process (D. Hunter, pers. comm.). A mycoplasma isolated from dead and live captive bighorn sheep with pneumonia was typed as M. arginini (Woolf et al. 1970). Apparently, no further work has been done regarding the pathogenicity of that strain to bighorn. With continued exposure of bighorn to domestic livestock, Mycoplasma spp. could interfere with recovery efforts. An outbreak of pneumonia associated with M. ovipneumoniae infection occurred in captive Dall's sheep (Ovis dalli dalli) following indirect exposure to domestic sheep (Black et al. 1988). Although Dall's sheep are unlikely to be exposed to domestic livestock in their native habitat, it is clear that captive herds or those in transit must be protected from exposure to pathogens.

Behymer et al. (1989) detected antibodies to Mycoplasma spp. in mule deer (Odocoileus hemionus) in California, but there was no association with disease, and isolations were not attempted. One-humped camels (Camelus dromedarius) and African buffalo (Syncerus caffer) on a game farm in Kenya had antibodies against Mycoplasma strain F38, and camels were seropositive for M. mycoides mycoides, but no isolations were made and no disease was seen (Paling et al. 1988). Kirchoffet al. (1996) found, in a survey of captive arthritic elephants (Elephas maximus and Loxodonta africana), that about 60% of the females harbored a new mycoplasma in the urogenital tract, although none was recovered from males. It is not known if the mycoplasma, named M. elephantis, causes arthritis.






18

Chronic Manifestations of Mycoplasmal Infections


Many mycoplasmal diseases are characterized by an overaggressive or

inappropriate immune response by the host, eventually leading to autoimmune damage to the affected sites, whether respiratory or urogenital tract, joints, heart, skin, or other organ systems (Krause and Taylor-Robinson 1992, Simecka et al. 1992, Cole 1996). Infected hosts may be more susceptible to secondary infections with other bacteria or viruses. In wildlife species, such complications may reduce the fitness of the animals by altering behavior, leading to decreased foraging efficiency, increased susceptibility to predators, or diminished mate seeking behavior. Energy that normally would be allocated to reproduction may be needed to repair or compensate for damage to multiple organ systems. Therefore, even if mycoplasmal infections do not cause mortality directly, they can affect individual and population viability.


Project Overview and Specific Objectives


Due to the listing by the Florida Game and Fresh Water Fish Commission of the gopher tortoise as a species of special concern, and the subsequent permitting of over 450 relocations involving more than 8000 tortoises, particular attention has been focused on the dynamics and persistence of both natural and relocated populations. Understanding the effects of URTD on individuals and populations is essential for proper management of remaining populations; therefore, a study was begun in 1993 on the etiology, pathology, and diagnosis of URTD in gopher tortoises. The original objectives were 1) to describe the pathology of natural infections and identify possible etiologic agents, 2) to perform






19


experimental infections to determine conclusively the etiologic agent and dose-related effects, 3) to refine the PCR test and delineate further the taxonomy of the tortoise mycoplasmas, 4) to characterize the immune responses of gopher tortoises to URTD pathogens, and 5) to compare the efficacy of culture, PCR and serology in detecting new infections.

As the original experiments were conducted and concern over the continued wellbeing of tortoise populations throughout the range increased, additional questions were raised. Those were concerned primarily with management and conservation of gopher tortoises, particularly with respect to relocation, repatriation, and restocking efforts. Several questions were raised repeatedly in meetings with state and federal agency personnel, particularly regarding possible immunity to the organism and factors related to transmission. In order to address those questions additional objectives were established, as delineated in specific objectives 2-7 below, and discussed in Chapters 3 through 8. Chapter 2 will detail the methods common to fulfilling the objectives.

This dissertation addresses the following specific objectives:

1) To describe the clinical presentation and lesions of natural infections by M. agassizii in

gopher tortoises and compare the lesions to those found in desert tortoises (Chapter 3). 2) To test the hypothesis that tortoises that have produced antibodies against M. agassizii

are protected against reinfection with the organism and subsequent development of

URTD (Chapter 4).

3) To assess the probability of horizontal transmission of M. agassizii between adult

tortoises and to test the hypothesis that transmission is more likely to occur from an






20

infected host that is clinically ill and PCR or culture positive than from a seropositive host that is not clinically ill or is clinically ill but culture or PCR negative (Chapter 5). 4) To determine ifM. agassizii is transmitted vertically (Chapter 6). 5) To assess the relationship of antibodies in eggs and hatchling serum to those in

maternal serum (Chapter 6).

6) To collect preliminary data to test the hypothesis that M. agassizii can be transmitted

environmentally (e.g., in burrows) (Chapter 7).

7) To discuss the implications of the findings of the above research and additional

concurrent research on conservation and management of gopher and other tortoises

(Chapter 8).















CHAPTER 2
METHODS



Tortoises, Intake Procedures, Clinical Assessments and Sampling Methods


For the natural infection studies, tortoises were obtained from various locations in the state of Florida under Florida Game and Fresh Water Fish Commission permits number WX93227 issued to Elliott R. Jacobson and number WX94037 issued to Mary B. Brown. Tortoises were processed the day of arrival at the University of Florida (UF), Gainesville. For the natural and experimental infection studies, gopher tortoises were transferred from a development site in central Florida to UF in April, July, and August 1994 and April 1995, and processed the day following arrival. Tortoises were examined for clinical signs of URTD: nasal and ocular discharge, palpebral edema, and conjunctivitis. The signs were graded individually on a scale of 0-3, none to severe. Total clinical sign score was calculated as the nasal discharge score plus the mean of three ocular sign scores (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. Straight line carapace length, thickness, and width were measured to the nearest cm with forestry calipers.





21







22

A 2-3 ml blood sample was drawn from the jugular or brachial vein and placed in a lithium heparin Vacutainer tube (Becton Dickinson and Company, Rutherford, New Jersey). Blood was centrifuged and an aliquot of plasma was removed for antibody screening by an enzyme-linked immunosorbent assay (ELISA) (see below).

After cleansing the area around the nares with alcohol dampened gauze, nasal flush samples were collected by flushing with approximately 0.5 ml sterile SP4 broth using a 1 ml syringe without needle. Calcium alginate-tipped swabs were inserted gently into the nares, and a sample was obtained and streaked onto SP4 agar plates.


Culture Procedures


A 100 pll aliquot of the flush sample was used for polymerase chain reaction (PCR) analysis; the remaining sample was serially diluted ten-fold to 10-2 and incubated at 300C for a maximum of three weeks, or until determined to be positive or contaminated. Twenty jpl of each dilution were placed on SP4 agar and incubated at 300C and 5% CO, as were the streak plates. Plates were examined regularly for a maximum of six weeks to detect the growth of mycoplasma. In the second year of the study, broth cultures were incubated for 24-48 hr before removing the aliquot for PCR, as that modification increased the sensitivity of the PCR (G. S. McLaughlin and D. R. Brown, unpub. data).


PCR Procedure


Nasal aspirate samples were analyzed for the presence of Mycoplasma agassizii DNA based upon PCR amplification of the 16S rRNA gene (Brown et al. 1995). Nasal flush and culture samples were centrifuged for 60 min at 14,000 rpm at 4C, and the







23

supernatant aspirated. Three to four microliters of 20 rpg/ml proteinase K (Sigma, St. Louis, MO) in 20 pi lysis buffer (100 mM tris pH 7.5, 6.5 mM DTT, 0.05% Tween 20) were added to the pellets, which were resuspended, and the samples were incubated at 370C for 8-16 hours. After denaturing the proteinase K at 970C for 15 min, 5 p of each sample were removed and added to 45 .pl of reaction solution containing two primers for the 16S rRNA gene at 1 ptM each, deoxynucleoside triphosphates at 200 jpM, 2.0 mM MgC1, and 2.5 units of Taq polymerase (Promega, Madison, WI). 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', Brown et al. 1995) and a Mycoplasma genus-specific region [anti-sense strand nt 1055 to 1031, 5'-TGCACCATCTGTCACTCTGTTAACCTC-3', Van Kuppeveld 1992]. Samples were subjected to 50 cycles of template denaturation for 45 sec at 94oC, primer annealing for 1 min at 550C, and polymerization for 45 sec at 72oC, followed by 10 min at 720C. Positive samples yielded 576 base pair (bp) products that were visualized by combining 15 gl of product with 2 pl bromphenol blue in 50% glycerol solution and electrophoresing on ethidium bromide-stained 1.5% agarose gels in tris-borate-EDTA buffer. Positive control samples using 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, Hind III digest of fX phage DNA, was included on each gel.

In order to confirm that the isolates obtained from naturally and experimentally infected tortoises were M. agassizii, an additional procedure, restriction fragment length polymorphism (RFLP) analysis, was conducted on at least one isolate from each tortoise. Twenty microliter samples of products from the above amplification procedure were







24

incubated with 10-20 units of the endonuclease Agel (New England Biolabs, Inc., Beverly, MA), which cuts the M. agassizii amplification product at nt 613, and 5 pl of reaction buffer, at 250C for one hour, and the products electrophoresed as above. The procedure resulted in products of 434 and 142 bp from M. agassizii-positive samples, and no change in non-M. agassizii-samples.


ELISA Procedure


An aliquot of plasma from each sample was used for determinations of levels of antibodies specific for M. agassizii (Schumacher et al. 1993). Ninety-six-well microtiter plates (Maxisorp F96, Nunc, Kamstrup, Denmark) were coated with 50 Pl of a whole-cell lysate ofM. agassizii strain 723 at 10pg/ml in phosphate buffered saline with 0.02% azide (PBS-AZ). Plates were incubated overnight at 4oC, 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 4C with 250 pl/well PBST containing 5% non-fat dry milk (PBS-TM). Following washing, 50 l of plasma diluted appropriately for the specific study with PBS-TM were added to individual wells in duplicate or triplicate, and the plates were incubated at room temperature for 60 min. The plates were washed, 50 pl/well of a biotinylated monoclonal antibody (MAb HL673) against the light chain of desert tortoise immunoglobulins IgY and IgM at 1 Pg/ml in PBS-TM was added, and plates were incubated for 60 min. Following washing, a conjugate of alkaline phosphatase and streptavidin (AP-S; Zymed Laboratories, Inc., San Francisco, CA) at 1:2000 in PBSAZ was added at 50 pl/well, plates were incubated for 60 min, and washed. Substrate, pnitrophenyl phosphate disodium (pNPP; Sigma), was prepared at 1 mg/ml in 0.01 M







25

sodium bicarbonate, pH 9.6, with 2 mM MgC12, and added to wells at 100 pl/well. Plates were incubated for 60 min in the dark, then read at 405 nm on a microplate reader (EAR 400 AT, SLT Labinstruments, Salzburg, Austria). The mean of two or three wells coated with antigen and incubated with conjugate and substrate only was used as the blank. A positive control, plasma from a naturally infected gopher tortoise from Sanibel Island, and a negative control, plasma from an uninfected tortoise from Orange County, were included on each plate.

Results from the ELISA were optical density (OD) readings from the microplate reader. The OD readings reflected the intensity of the yellow color developed when all components of the reaction (specific tortoise antibodies against M. agassizii, biotinylated MAb HL673, AP-S, and pNPP) were present. The readings are on a continuous scale, but were interpreted categorically by calculating the ratio of the sample readings to the negative control reading. Ratio values less than or equal to 2.0 were considered negative, those greater than 2.0 and less than or equal to 3.0 were classified as suspect, and those greater than 3.0 were classified as positive.



Study Group Assignment


Tortoises exhibiting one or more signs of disease, or with a positive culture or PCR result, were designated as diseased. Animals were designated clinically healthy if found free of signs of URTD, and with negative culture and PCR results. Tortoises were assigned to study groups based on sex, weight, clinical assessments, culture, PCR and ELISA results, and placed in the appropriate pens.







26

Husbandry Procedures


Tortoises were housed singly or in pairs in outdoor pens at the UF Animal

Resource Farm (ARF). There were four groups of 10 pens in a larger, chain link-fenced enclosure (Fig. 2-1). Pens were approximately 21 m2, constructed of a wooden frame with sheet metal extending vertically approximately 0.7 m above and below ground, partially covered by shade cloth, and provided with an artificial burrow, a water dish, and a cement feeding stone. Pens were observed daily by ARF staff and watered daily in the summer and as needed in the winter. The tortoises were fed a salad of mixed vegetables three times per wk, and fruit was provided on an occasional basis. In addition, I observed the pens and tortoises from three to seven days each week, with some days including multiple observations. Because of individual behavior patterns, not every tortoise was observed at each time point. The amount of food eaten was recorded for each pen the day following feeding, and the stones were cleaned. Remaining food was collected and bagged separately for each section, using brushes and dust pans assigned to that section. Stones were then sprayed with a dilute (1:30) bleach solution, allowed to soak for a few minutes, and hosed off. Water dishes were rinsed and filled daily, and bleached and scrubbed as necessary. 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 a different tortoise. Used gloves were placed in plastic garbage bags and disposed of properly. Before entering the first pen on a given day, personnel sprayed their footwear with a dilute bleach solution. Footwear was sprayed with bleach before leaving







27

a pen. Animals were captured by hand or using wire cage-type traps (Tomahawk Live Trap Company, Tomahawk, WI) that were covered with brown paper to protect the animals from the weather. Traps were cleaned, sprayed with bleach solution, and allowed to air dry following each use. 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, WI) for transport and holding. Containers were bleached, scrubbed, and washed in an automatic cage washer before reuse.










D C


A B







Figure 2-1. Layout of tortoise pens at the University of Florida Animal Resource Farm The outer fence was chain link, and the inner fence and pen dividers were corrugated sheet metal on a wooden frame.



Necropsy Procedures

All diseased and selected healthy tortoises were euthanatized with a combination of drugs. Ketamine was administered intramuscularly at 60 - 80 mg/kg followed by a







28

concentrated barbiturate solution (Socumb, The Butler Company, Columbus, Ohio, USA) intraperitoneally at I ml/kg. Once the tortoises showed complete muscle relaxation and were unresponsive to painful stimulation, they were exsanguinated via a 23 gauge butterfly catheter inserted into the carotid artery and then decapitated. Flush and swab samples were collected as previously described, then the head was bisected longitudinally with an electric saw. Following bisection, the cartilage over each nasal cavity was reflected aseptically, and flushes and swabs of 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 and the following tissues were collected, fixed in neutral buffered 10% formalin, sectioned at 5-6 pm, and stained with hematoxylin and eosin: glottis, cranial trachea, tracheal bifurcation, left lung, right lung, thyroid, heart, brain, thymus, esophagus, stomach, small intestine, pancreas, large intestine, cloaca, spleen, liver, left and right kidney, bladder, right and left gonads, chin gland, buccal salivary gland, chin gland, and tongue. Tissues were examined by light microscopy and abnormalities or changes were recorded.



Histopathology Procedures


For histopathologic studies, heads were fixed in 10% neutral buffered formalin

(NBF), decalcified, embedded in paraffin, sectioned longitudinally at 5-6 pm, and stained with hematoxylin and eosin. Sections were examined by light microscopy and classified on a scale of 0 to 5, with 0 being normal and 5 exhibiting severe inflammation and / or changes. Changes in the epithelium and submucosa were recorded separately.







29

The following criteria were utilized for grading lesions:

Normal (score = 0): Occasional small subepithelial lymphoid aggregates; rare heterophils in the lamina propria. No changes in mucosal or glandular epithelium; no edema.

Mild (1): Multifocal small subepithelial lymphoid aggregates; multifocally, small

numbers of heterophils, lymphocytes, and plasma cells in the lamina propria; mild edema in lamina propria; minimal changes in mucosal epithelium.

Moderate (2-3): 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; proliferation and disorganization of the basal epithelium.

Severe (4-5): Focally extensive to diffuse band of lymphocytes and plasma cells subjacent to and obscuring the overlying mucosal epithelium; large numbers of heterophils in lamina propria and infiltrating 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; occasional squamous metaplasia.


Statistical Analyses


Statistical analyses were performed using SAS (SAS Institute, 1988) or SigmaStat for Windows, Version 1.0 (Fox et al. 1994). Because the analyses varied for each experiment, the specific methods will be addressed in the appropriate chapters.















CHAPTER 3
NATURALLY OCCURRING UPPER RESPIRATORY TRACT DISEASE Methods


Tortoises

Twenty-three gopher tortoises from the following locations in Florida (Figure 3-1) were transported to UF from August 1993 to September 1995: Alachua County (n = 2), Sanibel Island, Lee County (n = 3), Volusia County (n = 1), St. Lucie County (n = 1), Indian River County (n = 1), Orange and/or Osceola Counties (n = 15). Collection of tortoises, except those from Orange and Osceola Counties, was opportunistic, and a result of submissions to the UF Wildlife Clinic, or other veterinary clinics. Some tortoises had exhibited signs of URTD, while others had been hit by automobiles. Tortoises from Orange and Osceola Counties were selected on the basis of clinical evaluations, ELISA, culture, and / or PCR results. Six clinically healthy animals were included in the latter group. Tortoises were evaluated for clinical signs of URTD (i.e., nasal and ocular discharge, palpebral edema, and conjunctivitis) and those exhibiting one or more signs of disease, with a past history of clinical signs, with positive culture or PCR results, or with a positive ELISA result, were designated as diseased. Tortoises were designated healthy if they were free of any history of or current signs of URTD and were culture, PCR, and ELISA negative.




30






31







2*
30

4*








Figure 3-1. Locations in Florida from which gopher tortoises were obtained. 1 - Alachua
County, 2 - Volusia County, 3 - Orange/Osceola Counties, 4 - Indian River
County, 5 - St. Lucie County, 6 - Sanibel Island, Lee County.



Necropsy and Histology Procedures

Necropsy procedure and light microscopic evaluation of tissues were performed as detailed in Chapter 2. Gross necropsies were conducted on 21 tortoises. Multiple tissues were collected from 15 tortoises, heads and livers from six , and only heads from two. Microbial Isolation

Flush and swab samples for Mycoplasma isolation were collected from the nasal passages and cavities of each tortoise and processed as described in Chapter 2. Swab specimens of the dorsal nasal cavities of 16 tortoises were collected for aerobic bacteria isolation and submitted to the Clinical Pathology Laboratory (CPL) of the College of Veterinary Medicine (CVM) at UF. Samples were cultured on Columbia blood agar and MacConkey's agar, and incubated at 370C. Bacteria were identified utilizing the







32


identification systems API 20E for enteric organisms and API NFT for non-enterics (BioMerieux Vitek, Hazelwood, MO, USA). Isolates of organisms consistent with Pasteurella were identified to species according to biochemical profiles listed for P. testudinis (Snipes and Biberstein 1982).


Electron Microscopy

Selected specimens were submitted to H. P. Adams of New Mexico State

University, Las Cruces, for scanning and transmission electron microscopic preparation and evaluation.

The left half of the bisected head of one healthy tortoise was prepared for

ultrastructural evaluations. The nasal cavity was instilled with 2.5% glutaraldehyde in 0.1 M phosphate buffer, then dissected out in its entirety, and selected areas were sampled. The tissues were dehydrated in an ascending series of ethanols and transferred to hexamethyldilisilazane for the final drying. The samples were sputter coated with gold and viewed by scanning electron microscopy (SEM).

Samples from 11 diseased tortoises were collected for transmission electron microscopy (TEM). The nasal cavity tissue was removed from the underlying cartilaginous tissues and separated into anterior dorsal, anterior ventral, posterior dorsal and posterior ventral quadrants. Each quadrant was cut into 1 mm cubes and placed in

2.5% glutaraldehyde, and post-fixed in osmium tetroxide. Specimens were prepared for TEM by embedding in epon-araldite and sectioning with an ultramicrotome. Thick sections were stained with toluidine blue and examined by light microscopy. Ultrathin sections were placed on copper grids, stained with uranyl acetate and lead citrate, and examined with a Hitachi H7000 transmission electron microscope.







33


Results


Based on clinical evaluations and diagnostic tests, eight tortoises were classified as healthy and 15 as affected by URTD. Infected tortoises were from Orange / Osceola Counties (n = 10), Lee County (n = 3), Indian River County (n = 1) and St. Lucie County (n = 1). Histological findings for the two groups will be discussed separately.


Normal Anatomy and Histology

The external nares opened into ventro-lateral depression, and were continuous with large dorsal nasal cavities (Figure 3-2). Right and left dorsal nasal cavities were separated by a cartilaginous septum. Each nasal cavity was bisected by a ridge, forming anterior and posterior compartments. Ventrally, the nasal passageways were continuous with the choanae (internal nares), which opened into the palatine region of the dorsal oral cavity.

The integument continued through the external nares into a short vestibule, which was initially lined by keratinized stratified squamous epithelium. That epithelium abruptly changed to mucous glandular epithelium, which lined the nasal passageway throughout its length. Interspersed among the mucous epithelial cells were ciliated epithelial cells. The ventro-lateral depression was lined primarily by mucous and ciliated epithelial cells (Figure 3-3a). Both anterior and posterior dorsal nasal chambers were lined by a multilayered olfactory epithelium with occasional mucous cells (Figure 3-3b). Numerous serous and mucous glands, vessels, nerve bundles, and clusters of melanophores were present in the connective tissue surrounding the nasal cavities. Small focal aggregates of lymphoid cells were seen in the submucosa.







34























' -'"7.. - ,.-.._._- -- -------Figure 3-2. Diagrammatic representation of the interior of a gopher tortoise head
sectioned longitudinally, illustrating the relationship of the nasal cavity to the
external and internal nares. Approximately 3x life size. Drawing by L.
Mallory.






35



a



















b



















Figure 3-3. Photomicrographs of normal gopher tortoise upper respiratory tract tissues.
Hematoxylin and eosin staining, 320x. a) Photomicrograph of the ventrolateral depression demonstrating the ciliated mucous epithelium; b)
Photomicrograph of the dorsal nasal cavity demonstrating the multilayered
olfactory epithelium. Photographs by E. R. Jacobson.






36


The bilateral thymus glands were difficult to find in healthy tortoises and were located cranial to the base of the heart, at the branching of the subclavian and carotid arteries. Grossly, the thymus was multilobulated. Histologically, there was a typically dark staining cortex and a lighter staining medulla. The cortex contained densely packed thymocytes. In the medulla there were significantly fewer cells including thymocytes, as well as thymic epithelial cells, myoid cells, and heterophils.

The spleen was located on the right side, between the proximal duodenum and

transverse colon and was associated closely with pancreatic tissue. Histologically, spleens were composed of distinct areas of white and red pulp. White pulp consisted of collections of lymphoid tissue surrounding blood vessels. Red pulp, located between the perivascular collections of the white pulp, included red blood cells within sinusoids and small numbers of lymphocytes.

The thyroid was located at the base of the heart. In one of the healthy tortoises, the thyroid was enlarged, with multifocal areas of follicular epithelial cell hyperplasia. In all tortoises, follicles varied in size, with many having numerous red blood cells in the colloid and either intraepithelial or supra-epithelial vacuoles.

Multiple foreign bodies were seen in the submucosa of the glottis of two tortoises, in the tongue of one tortoise, and in the buccal salivary gland of one tortoise. The foreign bodies were consistent with plant material. Lymphoid aggregates were scattered throughout the esophagus, small intestine, large intestine and cloaca, and also were present in the connective tissue surrounding the mental (chin) glands.

Of the healthy tortoises that were examined fully at necropsy, three were males and three were females. In one male, multifocal areas of mineralization of seminiferous tubules






37


were seen. Golden brown granules were scattered throughout the kidney within renal tubular epithelial cells of all tortoises. Mononuclear cells containing similar appearing granules were within the renal interstitium and the interstitium of the testes.


Pathologic Findings

Gross examination of heads of diseased tortoises revealed minimal to large

amounts of exudate within the nasal cavity and nasal passageways. Histologically, of the 15 heads, one had no changes, two had mild changes, seven had moderate changes, and five had severe inflammatory changes. In the tortoises with minimal changes, mild mucosal hyperplasia and slightly increased lymphoid aggregates were seen in the nasal passage and ventro-lateral depression. In those tortoises with moderate changes, the olfactory epithelia were usually normal, with only focal or mild changes in the submucosa. Changes generally were confined to the nasal passage and the ventral aspects of the nasal cavities, and consisted of mucosal epithelial and lymphoid hyperplasia, with infiltration of mononuclear cells and heterophils. In some tortoises with moderate inflammation, basal cell proliferation and loss of cilia could be detected. In the tortoises with severe inflammatory changes, there were lymphoid aggregates around submucosal glands, with glandular epithelial hyperplasia. The normal mucosal architecture was replaced by infiltrates of mononuclear cells and heterophils. The olfactory mucosa was replaced with proliferating mucous epithelial cells (Figure 3-4). Proliferating basal cells projected into the underlying lamina propria of some tortoises. Exudate, consisting of sloughed epithelial cells and inflammatory cells, was found in the nasal cavity lumen.

Infected tortoises had larger lymphoid aggregates in the submucosa of the glottis; several tortoises had focal areas of epithelial cell proliferation. Basal cell proliferation and







38

























id


















Figure 3-4. Photomicrograph of the nasal cavity tissues of a gopher tortoise with upper
respiratory tract disease. The changes were classified as severe, with
aggregates of lymphoid cells in the submucosa, proliferation of the basal cells,
and dysplasia of the mucosal epithelium. Hematoxylin and eosin staining,
320x. Photograph by E. R. Jacobson.






39


submucosal lymphoid hyperplasia were seen in the glottis of one tortoise. The hyperplasia extended into the cranial tracheal epithelium. In that tortoise, there also were multifocal areas of epithelial cell hyperplasia in the lung. Five other diseased tortoises had focal to multifocal lymphoid aggregates in the lung interstitia.

The gastrointestinal tracts tended to have increased numbers and larger lymphoid aggregates in the submucosa compared to those of clinically healthy tortoises. In one tortoise there was mucous cell hyperplasia of the colon and in another there was a severe colitis with basal epithelial cell hyperplasia and submucosal lymphoid hyperplasia, with infiltrates of large numbers of heterophils. Four other tortoises had increased lymphoid aggregates in the esophagus, stomach and/or small intestine. Three additional tortoises had increased lymphoid aggregates in the submucosa of the cloaca.

The kidneys of all diseased tortoises contained golden brown pigment granules

within renal epithelial cells. Hepatocytes of most tortoises contained similar granules, and pathologic changes were seen in the livers of nine tortoises. In seven, there were increased numbers and size of melanomacrophages in the liver and increased amounts of golden brown granules. One tortoise had cuffing of the central vein by lymphocytes and heterophils; another had aggregates of lymphocytes and melanomacrophages.

Electron micrographs of tissues of one tortoise from Sanibel Island and the tortoise from Indian River County demonstrated organisms consistent with Mycoplasma on the surface of the nasal mucosa (Figure 3-5). Associated epithelial cells had vacuolated cytoplasm and inflammatory cell infiltrates were present in the mucosa. Increased numbers of mucous epithelial cells were seen, consistent with light microscopic findings.






40

ELISA and PCR Results

ELISA and PCR results for both healthy and diseased tortoises are presented in

Table 3-1. All healthy gopher tortoises were seronegative for antibody specific against M. agassizii. Twelve diseased tortoises were seropositive, two were suspect, and one was seronegative. All healthy tortoises were PCR negative while 11 diseased tortoises were PCR-positive for Mycoplasma in nasal aspirates. Four clinically healthy tortoises that had negative culture and PCR results from nasal passage flush and swab samples had positive cultures and / or PCR results for samples from the nasal cavities. Microbial Isolation Results

The results of Mycoplasma and aerobic microbial cultures of the upper respiratory tract (URT) are presented in Tables 3-1 and 3-2. Mycoplasma was not cultured from the URT of any healthy tortoise. Mycoplasma was cultured from the URT of 11 tortoises with URTD. The aerobic microbial isolates of healthy tortoises consisted primarily of members of the genera Staphylococcus, Streptococcus, and Corynebacterium; a few Gram-negative rods were isolated. A greater number of Gram-negative species were isolated from the nasal cavities of tortoises with URTD, and those isolates made up a greater proportion of the isolates. Pasteurella testudinis was isolated from five diseased tortoises; in two it represented the major aerobic isolate.



Discussion


Gopher tortoises with clinical signs of URTD and evidence of exposure to

Mycoplasma agassizii were obtained from multiple sites in Florida. The light microscopic







41














































Figure 3-5. Transmission electron photomicrograph of the nasal cavity mucosa of a
gopher tortoise with upper respiratory tract disease. Organisms consistent
with Mycoplasma (arrow) can be seen in close association with host cell
membranes. Magnification 18,000x. Photograph by H. P. Adams.







42


Table 3-1. Summary of ELISA, PCR, culture, and nasal histopathology results from necropsied gopher tortoises from various locations in Florida, for determining infection with Mycoplasma agassizii.


Group Clinical ELISA PCR Mycoplasma Histopathology Signs Culture

Normal 0% 0% 0% 0% 0%
(n = 8)


Natural Infection 60% 85% 92% 80% 92% (n = 15)







Table 3-2. Summary of aerobic culture results from nasal cavity swabs of gopher tortoises from Florida.
Percent
Group Growth Species of isolates

Normal very scant to Staphylococcus spp. 45 (n = 5) moderate Gram-negative rods 25 a-hemolytic Streptococcus sp. 10 non-hemolytic Streptococcus spp. 10 Corynebacterium spp. 10


Natural very scant to Corynebacterium spp. 33 Infection heavy Pasteurella testudinis 18 (n = 11) Staphylococcus spp. 16 Gram-negative rods 15 Micrococcus sp. 11 Flavobacterium, Pseudomonas, 7 Lactobacillus, and others






43


changes in the upper respiratory tract of the gopher tortoises were similar to inflammatory and dysplastic changes reported for desert tortoises with URTD (Jacobson et al. 1991, Brown et al. 1994, Jacobson et al. 1995). However, inflammationa and epithelial proliferation around the glottis, tracheitis, and proliferative pneumonia have not been seen in desert tortoises. Additionally, two diseased tortoises had proliferation of the colonic mucosal epithelium. Those changes have not been seen in desert tortoises with URTD.

By electron microscopy, organisms consistent with Mycoplasma were

demonstrated on the nasal mucosal surfaces of two tortoises. Other than bacteria, no infectious agents were demonstrated in or on nasal cavity mucosa by electron microscopy. Eleven diseased gopher tortoises were PCR positive and M. agassizii was cultured from the upper respiratory tract of 11 diseased tortoises examined. These results support the hypothesis that M. agassizii is a cause of URTD in gopher tortoises in Florida.

The greater number of species and increased proportion of Gram-negative bacteria isolated from diseased tortoises could indicate that conditions in the upper respiratory tract of diseased tortoises are more favorable for the growth of those species, or that tortoises infected with M. agassizii are more susceptibile to opportunistic pathogens. The positive culture and / or PCR results from nasal cavity samples (obtained at necropsy) of four clinically healthy tortoises with negative nasal passage flushes and swabs support the hypothesis that tortoises can harbor the organism without showing clinical signs or shedding bacteria. Such animals may recrudesce under stressful conditions, begin shedding bacteria, and become infective to other tortoises.

Nine gopher tortoises exhibited pathologic changes in the livers, although the

significance of these changes is not understood currently. The mycoplasma may release







44


toxic compounds or induce the production of compounds by the tortoises that cause damage to the liver. Desert tortoises with URTD also show changes in liver tissue (B. L. Homer, pers. comm.), and some exhibit altered nitrogen metabolism (B. Henen, pers. comm.). The altered nitrogen metabolism may be due to behavioral changes leading to decreased foraging rates or efficiency, or it may be due to direct effects of the Mycoplasma infection, but the mechanism is not yet understood. Although only one of nine experimentally infected captive gopher tortoises significantly decreased its intake of vegetables (G. S. McLaughlin, unpub. data), and that decrease was temporary, wild tortoises may alter their behavior patterns to a greater degree. Alternatively, secondary infections by other bacteria may be the proximate cause of liver damage. However, no evidence of primary or secondary bacterial infection (i.e., necrosis) was seen histologically.

By the ELISA, none of eight clinically healthy tortoises and 11 of 15 diseased

tortoises had antibodies against M. agassizii, indicating previous exposure. Tortoises with negative ELISA results may have been in the early stages of the disease, when increased antibody levels had not occurred or were not detectable. The four diseased, seronegative, tortoises may have been infected with another agent. Westhouse et al. (1996) implicated an iridovirus as the cause of pneumonia, tracheitis, pharyngitis, and esophagitis in a gopher tortoise from Sanibel Island. The virus was readily detectable on both light and electron microscopy. It is not known if attempts were made to culture mycoplasma, or if an ELISA was run on a plasma sample. No indications of viral infections were seen in the tortoises examined for the current study.

Investigators recently have found seropositive gopher tortoises in Georgia (B.

Raphael, pers. comm.), seropositive, clinically ill, and/or PCR positive tortoises at a site in






45


northeast Florida, and seropositive tortoises at a site in Mississippi (D. M. Epperson, pers. comm.). Although many Florida and Georgia tortoise populations are fairly large, those in Mississippi are more restricted and are on the federal endangered species list. Outbreaks of URTD in populations with limited recruitment and no nearby source populations could contribute to severe declines in numbers and, possibly, local extinctions. Further studies, on a range-wide basis, need to be conducted to determine the distribution and potential impacts of URTD on gopher tortoise populations.

In addition, further research is necessary regarding the pathogenesis of URTD. While findings of liver, kidney, and intestinal tract lesions are interesting, they do not elucidate the mechanisms by which the changes are caused. Changes have been found in hormonal profiles of some infected desert tortoises (Rostal et al. 1996), which could lead to altered foraging and reproductive behavior, as well as decreased reproductive potential. If foraging behavior is affected, food and water intake might be reduced, which could affect liver and kidney functions. Both direct and indirect pathogenic mechanisms need to be studied in order to better predict the effects of URTD on tortoises.















CHAPTER 4
EFFECTS OF REPEATED EXPOSURE ON SEROPOSITIVE ADULTS



Introduction


In order to properly evaluate the results of the diagnostic tests and incorporate those findings into management and conservation plans, epidemiological questions, including the response of seropositive, asymptomatic tortoises to subsequent exposure to the agent, must be addressed. Although vaccines have been developed for some mycoplasmal diseases (Ellison et al. 1992, Lai et al. 1996, Markham et al. 1996), they are usually not fully protective (e.g., Djordjevic et al. 1996, Kleven et al 1996, Mohan et al. 1996, Washburn and Weaver 1996), and many natural infections by mycoplasmas do not engender a protective host immune response. The immune response is actually essential to the development of lesions, and infected animals are susceptible to repeated infection (Simecka et al. 1992). Due to the immunopathology, the disease may be more severe on subsequent exposure than on initial infection. I designed an experiment to test the hypothesis that tortoises that have produced antibodies against M. agassizii are protected against reinfection with the organism and subsequent development of URTD.











46






47


Methods


Acquisition of tortoises, intake and husbandry procedures, clinical assessments,

sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed in Chapter 2.


Statistical Analyses

The onset and severity of clinical signs of URTD, and ELISA, PCR, and culture results for the challenge tortoises were compared to those for control and naive tortoises using the SAS system (SAS Institute, 1988). Data from an additional naive animal infection experiment were included after it was determined that the data did not differ significantly from those collected from naive animals in this study. This ensured large enough sample sizes for meaningful comparisons at more time points. Differences in the severity of clinical signs, histologic lesions, and ELISA data among the three treatment groups were compared by an analyses of variance-type logistic regression using maximum likelihood estimators to compensate statistically for the different number of tortoises in each group at each sampling date. Percentages of tortoises showing clinical signs at different time points were compared by Fisher's Exact test, with a P value of 0.05 accepted as significant.


Experimental Design

Four groups of tortoises were established. Three groups [control (n = 6), naive (n = 11), and sentinel (n = 2)] had no history of exposure to M. agassizii, while the fourth, or challenge group (n = 8), had previous history of exposure as indicated by a positive result






48


on the ELISA. Initially, no tortoise in any group had clinical signs of URTD, or positive culture or PCR results. One tortoise originally slated for inclusion in the challenge group (i.e., seropositive, but clinically, culture, and PCR negative upon arrival) developed clinical signs before inoculation and was eliminated from the study.

Approximately 1 mo following arrival, the controls were sham inoculated

intranasally with 100 l sterile SP4 broth in each naris, and tortoises in the naive and challenge groups were inoculated in each naris with 100 gl of SP4 broth containing approximately 104 colony forming units (CFU) of M. agassizii strain 723, for a total dose of 108 CFU. The 723 isolate was obtained originally from a clinically ill tortoise from Sanibel Island, Lee County, Florida. The sentinel tortoises were captured, but received no other treatment.

Following inoculation, observations of all tortoises were attempted daily to

determine the onset and sequence of clinical signs. Behavior also was monitored. At 2 - 4 wk intervals post-inoculation (PI), tortoises were trapped, examined, and weighed, then tranquilized and blood, nasal flush and nasal swab samples were collected. Samples were processed as previously described. A total of 22 tortoises was examined at necropsy and histologically. Four control and nine naive tortoises were euthanatised and necropsied in October, 1994, before undergoing winter dormancy; seven challenge and two naive animals were euthanatised and necropsied in March, 1995, after emergence. Complete necropsies (n = 15) were performed on two control, six naive and seven challenge tortoises; only heads were examined on the other animals (n = 7).






49


Results


Clinical Signs

Of the six control group tortoises, none showed consistent clinical signs nor had positive culture, PCR, or ELISA results. At each of three time points, one control tortoise exhibited mild ocular signs that were probably associated with environmental conditions. One of two sentinels developed clinical signs at 12 wk PI, although she had not been inoculated with culture or medium. Of the naive tortoises, 67% exhibited clinical signs beginning 2-3 wk PI, 79% were clinically ill by 8 wk PI, and 94% had shown signs by 16 wk PI (Table 4-1). One naive tortoise never exhibited clinical signs. All seven challenged tortoises developed clinical signs of URTD before dormancy. Five (71%) challenged tortoises and one naive tortoise exhibited clinical signs soon after emergence. No tortoises became moribund or died during the study.

At 2 wk PI, six challenge and 17 naive tortoises were examined. Challenged

tortoises had higher total clinical sign scores than naive tortoises (2.6 vs. 0.5, P < 0.001), and higher scores for nasal discharge (1.5 vs. 0.1, P < 0.001), ocular discharge (0.8 vs. 0, P < 0.02), and palpebral edema (1.5 vs. 0.6, P < 0.02) (Table 4-1). Significantly more challenge than naive tortoises exhibited nasal and ocular discharge (67 vs. 6%, P < 0.01; 50 vs. 0%, P < 0.02; respectively) (Figure 4-1). No consistent differences were seen between the two groups at later sampling times (Figure 4-2).







50


Table 4-1. Percentages of naive and challenge tortoises positive for each clinical sign of URTD, and mean clinical sign scores. Scores <1.5 are classified as slight, >1.5 and <2.5 as moderate, and >2.5 as severe. ND = nasal discharge, OD = ocular discharge, ED = palpebral edema, CJ = conjunctivitis. P < 0.05 indicates significant difference between groups.

Time % Positive Mean Score
Sin Naive Challenge P Naive Challenge P
2 wk
Total 67 83 0.626 0.5 2.6 0.000 NDa 6 67 0.007 0.1 1.5 0.000 OD 0 50 0.011 0 0.8 0.015 ED 59 83 0.369 0.6 1.5 0.012 CJ 53 83 0.340 0.6 1.0 0.345
4 wk
Total 79 80 1.000 1.5 1.9 0.425 ND 50 60 1.000 0.8 1.2 0.254 OD 46 20 0.366 0.8 0.6 0.666 ED 52 40 1.000 0.7 0.8 0.864 CJ 59 60 1.000 1.0 0.8 0.670
6 wk
Total 94 100 1.000 2.1 2.7 0.229 ND 67 83 0.626 1.1 1.9 0.040 OD 44 57 0.673 0.5 1.0 0.118 ED 72 71 1.000 1.0 0.9 0.652 CJ 89 43 0.032 1.4 0.7 0.055
8 wk
Total 82 83 1.000 2.6 2.4 0.688 ND 71 71 1.000 1.4 1.6 0.677 OD 54 57 1.000 0.9 0.7 0.479 ED 71 71 1.000 1.1 1.3 0.634 CJ 71 29 0.075 1.5 0.6 0.004
12 wk
Total 83 83 1.000 2.4 2.4 0.989 ND 62 71 0.642 1.5 1.6 0.836 OD 54 57 0.676 1.0 0.7 0.418 ED 67 71 0.667 0.7 1.3 0.068 CJ 79 29 0.198 1.1 0.6 0.105
Nx
Total 84 83 1.000 2.6 2.4 0.744 ND 79 71 0.646 1.4 1.6 0.733 OD 79 57 0.340 1.2 0.7 0.123 ED 58 71 0.668 1.0 1.3 0.283 CJ 63 29 0.190 1.3 0.6 0.036









100

90 M Naive

. 80 -OChallenge

70 *


60

S 50 o 40

30

e 20 :

10



CS Total ND OD ED CJ

Figure 4-1. Percent of gopher tortoises infected with Mycoplasma agassizii positive for any clinical sign and each individual sign at 2
weeks postinfection. Tortoises in the challenge group had serological evidence of prior exposure to M. agassizzi, but
those in the naive group did not. CS = clinical signs, ND = nasal discharge, OD = ocular discharge, ED = palpebral
edema, CJ = conjunctivitis. * Indicates significant difference between groups, P < 0.05.









3.50

** EControl

3.00 - WNaive
OChallenge

2.50


. 2.00 *


I
S1.50


1.00


0.50


0.00 1 -
2 4 6 8 12 Nx Weeks Postinfection Figure 4-2. Total clinical sign scores for three groups of tortoises, one control and two experimentally infected with Mycoplasma agassizii.
Bars represent standard errors. ** Indicates a significant (P < 0.05) rise in clinical sign score and a significant difference
between groups. * Indicates a significant rise in clinical sign score.







53


Culture and PCR Results

The sentinel tortoise that became ill also had positive culture and PCR results. All inoculated tortoises had at least two positive culture results, and 17 of 18 had positive cultures from samples collected at necropsy. Mycoplasmal DNA was detected by PCR analysis from each inoculated tortoise, including those that did not exhibit clinical signs of URTD when sampled. By RFLP analysis, all isolates corresponded to M. agassizii. ELISA Results

There was no anti-M. agassizii antibody response by the control tortoises to sham inoculation, although one sentinel had seroconverted by 4 wk PI. All inoculated tortoises seroconverted or had significantly increased ELISA values. ELISA values were greater for challenge than naive tortoises at each time point (Table 4-2, Figure 4-3). A significant increase in ELISA values of challenged tortoises was observed by 4 wk PI (mean ratio of samples to negative control of 6.3 vs. 3.3 at 0 wk, P < 0.05), and seroconversion of naive tortoises was observed by 6 wk PI (ratio of 3.9 vs. 1.5 at 0 wk, P < 0.05). The ratio of the challenge to the naive ELISAs increased at 2 and 4 wk PI, then declined.



Table 4-2. Least-squares mean ELISA values for naive and challenge tortoises. P < 0.05 indicates a significant difference between groups.

Time Naive Challee P
0 weeks pi 0.2049 0.4707 0.0437 2 weeks pi 0.2235 0.5925 0.0126 4 weeks pi 0.2439 0.8848 0.0001 6 weeks pi 0.5464 1.5254 0.0001 8 weeks pi 0.8314 1.8219 0.0001 12 weeks pi 1.4062 1.9839 0.0001 Necropsy 1.6796 2.2889 0.0001









2.50


EControl
2.00 MNaive OChallenge
**


1.50 **



S1.00

* **

0.50



0.00
0 2 4 6 8 12 Nx Weeks Postinfection Figure 4-3. ELISA results for three groups of tortoises, one control and two experimentally infected with Mycoplasma agassizii.
Tortoises in the naive group had no previous exposure to M. agassizii, while those in the challenge group had evidence of prior exposure. Line indicates ratio value (sample to negative control) of 3.0. * Indicates significant differences (P < 0.05) between challenge and naive groups. ** Indicates significant differences between naive and control groups.
*** Indicates the first significant differences from Time 0 value for that group. Bars represent standard errors.





L-.0damm






55


Histology Results

The upper respiratory tracts of the four control tortoises examined at necropsy had normal histologic appearances. In contrast, all tortoises inoculated with mycoplasma showed lesions similar to those seen in naturally occurring URTD. All challenge tortoises had moderate to severe inflammation and changes in the epithelium and submucosa. Three naive tortoises had minimal lesions and eight had moderate to severe abnormalities.. Lesions were consistently seen in the ventrolateral depression of the nasal cavity, a region immediately caudal to the vestibule (see Chapter 3).



Discussion


No tortoises inoculated with sterile medium developed clinical signs or

histopathologic lesions, indicating that lesions were not a result of the mechanical effect of the medium on the tissues, nor of host inflammatory response to the medium. Experimentally infected naive animals did not seroconvert until 6 to 8 weeks PI. Because the sentinel that became ill and seroconverted was sampled only on arrival and 8 wk following arrival (4 wk PI for the inoculated tortoises), the timing of its seroconversion is uncertain. No known clinically ill tortoises were transported or held with the experimental animals once they left the Orange County holding facility, and the sentinel was not held in the facilities or with other tortoises during the time tortoises were being inoculated. Additionally, in an experiment involving the transfer of healthy tortoises into pens immediately following the removal of ill tortoises, there was no indication of environmental transfer, although the sample sizes were very small (see Chapter 7).






56


Therefore, the sentinel whose seroconversion was detected 8 wk after arrival probably was exposed to M. agassizii just before transport to UF.

Tortoises harboring M. agassizii may not show clinical signs, may exhibit mild signs, or may show signs only intermittently. Because tortoises not showing any ocular signs or nasal discharge sometimes have positive culture and PCR results, the organism may be transmissible from asymptomatic tortoises under appropriate conditions. Due to unavoidable constraints on sampling live animals, I cannot exclude the possibility that clinically healthy, seropositive, culture and PCR negative animals harbor the bacteria. As shown by the two tortoises (the sentinel and the tortoise that was eliminated from the study) that initially had negative culture and PCR results but developed disease without being inoculated experimentally, some infections may go through an extended latent period with low numbers of organisms in the nasal passages, or animals that appear to have cleared the organism may, in fact, have not.

The clinical response of challenged animals was more rapid and severe than that of naive animals, indicating that no protection was conferred by previous exposure to the organism. This is consistent with some other mycoplasmal diseases in which the immune response confers limited or no protection (Ellison et al. 1992), or contributes to pathogenesis, such as arthritis in fowl caused by M. synoviae (Kume et al. 1977), conjunctivitis in cattle caused by M. bovoculi (Rosenbusch 1987), and pneumonia in humans caused by M. pneumoniae (Krause and Taylor-Robinson 1992). Tully et al. (1995) have found that some mycoplasmal surface proteins share sequence and structural homologies with vertebrate proteins, and suggest that these may play a role in eliciting autoimmune responses. Repeated exposure to mycoplasmal proteins that resemble a






57


host's proteins may sensitize the host and induce an autoimmune response, leading to chronic manifestations of disease even if the primary etiologic agent is cleared.

Although actively produced antibodies apparently do not confer protection to adult tortoises, it is unknown if passively transferred antibodies (see Chapter 6) provide protection against URTD for hatchlings. Preliminary observations indicate that seronegative hatchlings are at least as susceptible to infection as adults, and that the disease progresses more rapidly, with high morbidity in the first 6 mo PI (see Chapter 5). Desert tortoise hatchlings exposed to infected adults developed clinical signs of URTD and suffered extremely high mortality rates (51 of 69, or 74%, of those with clinical signs died) (Oftedal et al. 1996). Mycoplasma agassizii was isolated from necropsied individuals, and severe inflammation was seen histologically. The immune status of the hatchlings and parents was unknown prior to the development of the outbreak, so no inferences can be drawn regarding the potential role of maternal antibodies in the course of URTD in hatchlings. Because cellular components probably play a major role in the pathogenesis of URTD and those are not transferred vertically in reptiles, the maternal antibodies may provide some protection. It would be interesting to explore the question of increased susceptibility to or severity of disease vs. protection from disease in hatchlings that have received maternal antibodies against M. agassizii.

The immune response stimulated by previous exposure to M. agassizii did not

prevent reinfection or ameliorate signs of URTD after subsequent inoculation. The onset of clinical signs was, in fact, more rapid and the initial severity was greater than in first infections. Development of specific antibodies against M. agassizii did not ensure clearance of the organism by the tortoises, either on initial or subsequent exposure. Based






58


on this study, tortoises testing positive for antibodies against M. agassizii cannot be considered good candidates for release in repatriation, restocking, or relocation efforts. It may be acceptable to release seropositive tortoises in areas that already have seropositive animlas, as long as the overall prevalence is not increased significantly. Seropositive tortoises may provide desirable genetic material (see Chapter 6), and would be valuable as research subjects to study interactions with other disease agents and the effects of repeated exposure on metabolism, autoimmune responses, reproduction, and survival.















CHAPTER 5
HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII



Introduction


In order to develop management and conservation plans that incorporate the

potential of disease to affect populations, epidemiologic questions must be addressed. Of particular concern are the probability of transmission of the disease organism from one tortoise to another (horizontal transmission), the rate of spread within populations, and the potential for spread to nearby populations. The probability of transmission may be related to a tortoise's clinical and culture or PCR status. Infected tortoises may be culture and/or PCR positive without showing clinical signs. Conversely, it is sometimes difficult to detect bacteria in clinically ill tortoises. I designed an experiment to test the hypothesis that horizontal transmission of M. agassizii will occur only from those individuals that are clinically ill, and PCR and/or culture positive.


Methods


Acquisition of tortoises, intake and husbandry procedures, clinical assessments,

sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed in Chapter 2.





59






60


Experimental Design

Fifteen pairs consisting of one male and one female were established in August, 1994. Due to the limited availability of seropositive and/or clinically ill tortoises, the sample design was not balanced. Five pairs of asymptomatic, ELISA-, PCR-, and culturenegative control tortoises were established as controls. The other ten pairs consisted of a clinically healthy, ELISA-, PCR-, and culture-negative tortoise that had been at UF since April, 1994 (resident), and an ELISA-positive or -suspect (irrespective of PCR or culture status) tortoise of the opposite sex and similar size (Figure 5-1, Table 5-1). One resident female was paired with an ELISA-negative, but clinically ill, culture- and PCR-positive male. The serosuspect and two seropositive tortoises were clinically healthy, and had negative culture and PCR results. The remaining six seropositive tortoises showed moderate clinical signs of illness. Three of those six were culture- and PCR-negative, one was culture- and PCR-positive, one was culture-positive but PCR-negative, and one was culture-negative and PCR-positive.

Behavioral observations and clinical signs of URTD were recorded

opportunistically (see Chapter 2), and blood and nasal flush samples were collected from all tortoises in August and October, 1994, and in March and August, 1995. In 1996, samples were collected in February or March, and in July or August. Clinical signs and weights were recorded, and photographs were taken at each sampling time. Data were analysed using Chi-square and Fisher Exact tests for differences in proportions of tortoises becoming infected under different exposure conditions. Infected animals were euthanatized in July, 1996, except as noted below, and uninfected tortoises were released pursuant to Florida Game and Fresh Water Fish Commission permit number WR96224.







61












Seronegative Clinically ll
Culture & PCR
Positive Clinically Healthy
n 1 Culture & PCR
Negative Culture & PCR Seropositive n 3 Negative
or Suspect Cnayn = 3
n 9 FClinically III
n =6 Culture & PCR Positive
n= 1


Culture or PCR Positive
n= 2




Figure 5-1. Flow chart showing initial distribution of seropositive, clinically ill, culture and PCR positive tortoises.










Table 5-1. Clinical signs, culture, PCR, and ELISA status of gopher tortoises included in the upper respiratory tract disease pairing study at each sampling time. CS - clinical signs (nasal or ocular discharge, palpebral edema, and/or conjunctivitis), Cl - culture results, P - PCR results, E - ELISA results.

Pen ID Aug. 1994 Oct. 1994 Mar. 1995 Aug. 1995 Feb.-Mar. 96 Jul.-Aug. 96 Necropsy No. Gender CS Cl P E CS C1 P E CS Cl P E CS Cl P E CS Cl P E CS P E Date Al 226 F - - - - - - - - - - - - - - - - - -- R
201M- --------------------- R
A2 577F-------------------- + -b 7/96
147M ------------+ + + -+ - + + + 7/96
B3 261F----------------------- R
160 M----------------------- R
B4 129 F - -211M----------------------- R
B5 108 F ----------------------- R
140 M ----------------------- R
D1 135F ------------++- s + + + + + 7/96
241M - - - - + - - -+ + + + + + + + + + 7/96 D2 275F + --+---+ + + - -++- -+ - - + + 7/96
144 M -- ----- disappeared U D3 151F + + + + + + s + + - + + - - s + - - - + s + 7/96
126M- - - - - + + + + - + + + + - + + - -+ - + 7/96 D4 186F- s + + --------- - + +-+++-+ + + 7/96
235MM + + + + + +- - + + - - + s -+ 7/96 D5 213F - - -+ - + + + + + + + + + euthanatised 6/95
185M + - ++ + + + + +- + + + - + euthanatised 6/95 D6 150F - - - + ++ s + + + + + s - + + + - +1+ - + 7/96
183M + + + - + + + + + + + + - + disappeared Uh









Table 5-1--continued-Pen ID Aug. 1994 Oct. 1994 Mar. 1995 Aug. 1995 Feb.-Mar. 96 JuL-Aug. 96 Necropsy No. Gender CS Cl P E CS C1 P E CS C1 P E CS Cl P E CS Cl P E CS P E Date D7 190 F - - - - s - - - s -----disappeared IU'
228M - - -+---+ - - -+ - - - + s -- ++ -+ + 7/96
D8 257F + - - + + - + + - + + - - + + + - + + - + 7/96 130M -------- s - - + + s + + -+ + + 7/96
D9 205F - --s+ --- +- - - + --- s - H
110M----------------------- R
D10 287 F + - + - + - - - + - + + euthanatised 6/95
125 M ---- s-s- +- - - s - - - s--- - - - R
"Released pursuant to Florida Game and Fresh Water Fish Commission permit number WR96224. bDenotes suspect status. Clinical signs: very mild, transient signs that could have been associated with environmental conditions or mechanical irritation. Culture: presumptive colony(ies) of Mycoplasma on agar that failed to grow when transferred to broth, or were overgrown by contamination before definite determination could be made. PCR: a very faint, nearly undetectable, signal. For ELISA, a ratio value between 2.0 and 3.0.
cBurrowed into an adjacent pen and was exposed to an experimentally infected tortoise. dHibernating as of December 1, 1996.
'Disappeared, presumed dead in burrow. Last examined August, 1995. fEuthanatised due to uterine rupture.
Euthanatised due to poor clinical prognosis.
hDisappeared, presumed dead in burrow. Last examined March, 1996. 'Disappeared, presumed dead in burrow. Last examined May, 1996. jUnable to evaluate ocular signs prior to administration of ketamine. kEuthanatised due to failure to reproduce and large mass viewed on radiography.






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Results


General Observations

Observations indicated varying degrees of interaction between pair members.

Several males were excluded from the original burrow in the pen by their female partners. Until one of the two dug a new burrow, those males, particularly 125, 185, 201, and 235, spent most of the day in a corner of the pen, under the shade cloth, and sometimes spent the night out of the burrow. Other pairs spent considerable time in courtship, or foraging in close proximity to one another.

In March, 1995, five hatching tortoises were found in pen DI. The female (135) may have been gravid or storing sperm when she arrived at the UF facility in April 1994, and laid the clutch during the summer of 1994. Alternatively, she may have copulated with the male (241) and ovulated shortly after the pairing in August 1994. Based on gopher tortoise physiology (Taylor 1982), however, I believe the former explanation is more likely. One hatchling was removed and euthanatised due to congenital abnormalities; the other four were left in the pen.

Three tortoises were euthanatised in June 1995 (Table 5-1). One, number 287 (pen D10), was an initially seropositive, clinically ill, culture and PCR negative female whose partner (125) had not become ill. She failed to reproduce, and a large mass could be seen on radiographs. At necropsy, a large urolith (bladder stone), approximately 5 cm in diameter, was removed. The second, number 213 (pen D5), was an initially seropositive, clinically ill, culture and PCR positive female whose partner had become ill within one month of pairing. She failed to lay six eggs, and on exploratory surgery it was discovered that both oviducts had ruptured and the eggs were free in the coelomic cavity.






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She was euthanatised without recovery from anesthesia. Her partner, number 185, was clinically ill, lethargic, and losing weight, and was euthanatised shortly thereafter.

One healthy, seronegative male tortoise (144, pen D2), paired with a seropositive female (275) disappeared after March 1995 (Table 5-1). He may have been overturned during interactions with the female and been pushed into the burrow, where he would have been unable to right himself. At the time of his disappearance, he had exhibited no indications that transmission of M. agassizii had occurred. The male tortoise from pen D6 (183) also disappeared, and may have met a fate similar to that of 144. His disappearance may have been related to agonistic interactions with any of the tortoises originally in pens D1, D6, D7, or D8, as discussed below. Due to the nature of tortoise burrows, it was impossible to determine unequivocally the fates of the animals that disappeared.

Approximately 10-11 months after pairing, in June and July 1995, the burrowing activities of the large number of tortoises in the small area resulted in the communication of several pens (Figure 5-2). Tortoises in pens D1, D6, D7, and D8 moved regularly between pens and had numerous agonistic encounters with one another. The four tortoises in pens D3 and D4 also moved between pens and had frequent interactions. Although tortoises from pens D9 and D10 also interacted with each other, aggressive interactions were not observed. One control male (147, pen A2), burrowed into an adjacent pen (A7) containing an experimentally infected ill tortoise, and moved between the two pens over several months. Neither the control female (577) nor the third tortoise was observed to switch pens. The study was originally designed to last for 12-14 mo, but when the animals began moving between pens, it was extended to 24 mo in order to collect more data.






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D10 287F D9 205F D8 257F D7 190F D6 150F 125M 110M 130M 228M 183M



D5 D4 D3 , D2 ! . 151F 275F
213F 186F 126M 135F J
185M 235M _ 144M 241M Figure 5-2. Burrow map showing interconnections among pens Dl, D6, D7, and D8; between D3 and D4, and between D9 and D10. * Burrow entrances.



Evidence of Transmission of Mycoplasma agassizii

The male control tortoise (147) became ill, seroconverted, and had positive culture and PCR results (Table 5-1). His partner became clinically ill and seroconverted after he returned to his original pen. No other control tortoises became ill or seroconverted. The D-section burrowing activities mentioned above resulted in the exposure of all tortoises in pens Dl and D7, which until that time had been clinically healthy, to clinically ill tortoises with positive culture and/or PCR results. Three became clinically ill, and had increased ELISA readings, with one of two initially negative tortoises seroconverting. Those activities also resulted in the exposure of the tortoises in pen D8 to clinically ill, culture and PCR positive tortoises. The seronegative male (130) became ill, had positive culture results, and seroconverted; the clinically ill, seropositive female (257), which had been culture and PCR negative until that time, also became culture positive. All four hatchlings in pen D1 became severely ill, seroconverted, and had positive culture results. All were euthanatised, and pure cultures of M. agassizii were recovered from the nasal cavities and conjunctiva on necropsy.






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The initially seronegative, clinically ill, culture and PCR positive male (183) had seroconverted by October 1994. Based on his original samples from July 1994, and the ELISA values of his October samples, he probably seroconverted within 2 wk of pairing, and was considered as ELISA positive for the analyses (see below). In the ten experimental pairs, eight initially asymptomatic members, two of which were originally ELISA-positive and six of which were originally ELISA-negative, became moderately ill with signs of URTD and had increased ELISA values, with the latter six seroconverting. Two other tortoises (125 and 190) showed mild clinical signs that may have been associated with mechanical irritation or environmental conditions, but did not seroconvert.

Of the four initially uninfected tortoises that showed no increase in antibody levels, three did not have positive culture or PCR results. One (190) had only mild ocular signs, and two (110 and 144) never showed clinical signs. The fourth (125), which consistently showed mild signs and had a suspect PCR result in October 1994, was exposed to a tortoise (287) that had positive samples only at necropsy. Based on observations, they had only limited interactions. The serosuspect female (205), who was in contact with both 110 and 125, showed occasional, transient, mild clinical signs, but never had positive cultures or PCR results, nor did her antibody levels increase. In fact at all sample times after August 1994, she was classified as seronegative.

Of the 16 originally culture and PCR negative tortoises, 10 later had positive or suspect culture or PCR results. Eight were exposed to clinically ill, culture or PCR positive tortoises, seroconverted and had positive results, the ninth had positive culture results only from necropsy samples, and the last was the individual (125) discussed above. No tortoise became ill unless exposed to another ill tortoise.






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Transmission Probabilities

Analysis of the data was complicated by the changing exposure status of the

tortoises due to the housing problems discussed above. Therefore, analyses were carried out on three data sets. The first consisted of data from the first 10 mo (August 1994 June 1995) of the study, the second from the last 14 mo (June 1995 - July 1996), and the third of the cumulative data. The first data set (Table 5-2) consisted of the original 15 pairs. The second set (Table 5-3) did not include those pairs in which transmission had already occurred, but only the control pairs and those whose exposure status had changed, leaving 11 observations. The third set (Table 5-4), with 22 observations, included some tortoises twice, as the observations of interest were the exposure events themselves, and not the pairs or individual tortoises. Although sample sizes were very small, and some cells had no observations, chi-squared tests of differences in proportions were run on the three data sets. The final set was collapsed on the three predictor variables (ELISA, clinical, and culture/PCR status; Table 5-5) to determine which had the most influence on transmission probability.

Four exposure status classes were established based on ELISA classification, clinical signs, and culture or PCR status. Class assignment was based on a tortoise's status at any point during the time period of interest. Therefore, a tortoise's status could change between the first and last parts of the study. Each criteria was recorded as positive or negative, and transmission was recorded as yes or no. Transmission was classified very conservatively, with any tortoise exhibiting even mild, transient, clinical signs not obviously related to mechanical irritation (e.g., plant material in the eye or nose) or environmental conditions (e.g., being held overnight in a box in which the tortoise had






69


urinated), or having one suspect culture or PCR result recorded as having been infected by horizontal transmission. The four classes were 1) ELISA, clinically, and culture/PCR negative (-/-/-), 2) ELISA positive, clinically and culture/PCR negative (+/-/-), 3) ELISA and clinically positive, and culture/PCR negative (+/+/-), and 4) ELISA, clinically, and culture/PCR positive (+/+/+). The first data set (Table 5-2) consisted of the following observations (class, yes, no): 1,0,5; 2,0,2; 3,2,2; and 4,4,0. The second set (Table 5-3), for the last 14 mo, was comprised of the following observations: 1,0,4; 2,0,1; 3,0,1; and 4,5,0. The last, cumulative, data set, included the following observations: 1,0,5; 2,0,3; 3,2,3; and 4,9,0. Chi-square tests of association of exposure status and transmission were significant for all data sets, with 3 degrees of freedom (df) and P < 0.001 for all sets (Tables 5-2, 5-3, 5-4).



Table 5-2. Contingency table for exposure status and transmission occurrence for the first 10 mo (August 1994 - June 1995) of the gopher tortoise URTD pairing study.

Exposure Status Transmission ELISA/Clinical/Culture or PCR Yes No
-/-/- 0 5 +/-/- 0 2 +/+/- 2 2 +/+/+ 4 0 , 3 df= 10.8, P = 0.013, power = 0.80 Table 5-3. Contingency table for exposure status and transmission occurrence for the last 14 mo (June 1995 - July 1996) of the gopher tortoise URTD pairing study.

Exposure Status Transmission ELISA/Clinical/Culture or PCR Yes No
-/-/- 0 4 +/-/- 0 1 +/+/- 0 1 +/+/+ 5 0 2, 3 df= 11.0, P = 0.012, power = 0.81






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Table 5-4. Contingency table for exposure status and transmission occurrence for the duration (August 1994 - June 1995) of the gopher tortoise URTD pairing study.

Exposure Status Transmission ELISA/Clinical/Culture or PCR Yes No
-/-/- 0 5 +/-/- 0 3 +/+/- 2 3 +/+/+ 9 0 2, 3 df= 17.2, P = 0.0006, power = 0.96




Table 5-5. Transmission probabilities and tests of significant differences of proportions for different exposure status classes of tortoises relative to upper respiratory tract disease.

Exposure Transmission Probability of Fisher Exact
Status Yes No Transmission Test, P value
ELISA
negative 0 5 0
positive 11 6 0.65 0.018

Clinical illness
negative 0 8 0
positive 11 3 0.79 0.001

Culture / PCR
negative 2 11 0.15
positive 9 0 1.00 0.0002



Because transmission was defined very conservatively for the purposes of this

study, two tortoises were categorized as having been infected via horizontal transmission

based solely on the appearance of mild clinical signs. Neither animal seroconverted nor

had positive or suspect culture or PCR results during the second half of the study. Both

tortoises were housed with clinically ill, seropositive tortoises that were culture and PCR

negative. If those tortoises were designated negative relative to transmission, the final






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data set was comprised of the following observations: 1,0,5; 2,0,3; 3,0,5; and 4,9,0. The chi-squared value was 22.0, with 3 df and P < 0.0001.

When Table 5-4 was collapsed on ELISA status (Table 5-5), none of the tortoises in five ELISA negative pairs became ill or seroconverted, but of the 17 interactions involving ELISA positive tortoises, 11 resulted in transmission, for a probability of 0.65. Fisher Exact test of a difference in proportions was significant at P = 0.018. When collapsed on clinical status, none of eight interactions involving negative tortoises resulted in transmission, as defined above, while 11 of 14 (probability 0.79) involving clinically ill tortoises did (Fisher exact test, P = 0.001). If transmission was defined more liberally, and designated as positive only if a tortoise seroconverted and had clearly positive culture or PCR results, then nine of 14 (probability 0.64) interactions with clinically ill tortoises resulted in transmission (Fisher exact test, P < 0.006). When transmission was defined conservatively, for culture and/or PCR status, 2 of 13 culture/PCR negative class interactions resulted in transmission (probability 0.15), while all of nine positive class pairings effected transmission (Fisher exact test, P = 0.0002). If transmission was defined more liberally, then none of 13 culture/PCR negative interactions resulted in transmission, and the Fisher exact test was significant with P = 2 x 10'.



Discussion


The unplanned movements of the tortoises resulted in more exposure events than initially planned, and possibly compromised some of the original experimental design. However, at least one pair remained in each exposure category and was followed for 24






72


mo, twice as long as originally planned. Therefore, the intent of the study was not compromised by the tortoises' natural behavior.

Although manifestation of latent infections could not be excluded absolutely, the results supported the hypothesis that M. agassizii was horizontally transmitted between adult tortoises, and between adult and hatchling tortoises. The route of transmission may have been direct contact, aerosol, or fomite transmission. Direct contact was the most likely route. If aerosol transmission occurred, it was probably over short distances, such as would be likely to lead to direct contact. Sentinel and control animals housed in pen groups that also housed ill tortoises did not become ill or seroconvert, indicating that aerosolized bacteria were unlikely to travel even relatively short differences or over low (0.5 m) barriers. Similarly, fomite transmission is unlikely to play a major role (see Chapter 7). However, because many tortoise interactions occur in burrows, it is difficult to assess the importance of the various modes.

Although sample sizes were small, results supported the hypothesis that

transmission is more likely when the "donor" is symptomatic, although tortoises without clinical signs may be infected and able to transmit the pathogen under appropriate conditions. After rainstorms, it is not uncommon to find two or more tortoises drinking from the same puddle (R. E. Ashton, pers. comm.; G.S.McLaughlin, pers. obs.). When tortoises drink, they often get water in their noses, which they then blow or "sneeze" out. An asymptomatic tortoise harboring bacteria may shed enough in this manner to infect a nearby conspecific via the aerosol route, or to leave an infective dose in the water or on nearby plants.






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A concurrent study (Brown et al. 1996b), in which gopher tortoises were

experimentally infected with varying doses of M. agassizii, demonstrated that the infective dose of the 723 strain is very low, less than 10 CFU. However, an additional isolate failed to induce disease or elicit an antibody response. In previous research involving desert tortoises, Brown et al. (1994) demonstrated that exudate from ill tortoises elicited a stronger immune response and induced more severe disease in inoculated tortoises than pure cultures of M. agassizii. In addition to the mycoplasma, the exudate probably contains other species of bacteria (e.g., Pasteurella testudinis, Corynebacterium sp.), as well as cellular components and cytokines that may elicit an immune response and contribute to the disease process.

The two tortoises that exhibited mild signs without seroconverting or having positive culture and PCR results may have been responding to cellular or chemical components in the exudate from their partners, to environmetnal conditions, or to other species or strains of bacteria that were only mildly pathogenic. Because conjunctival swabs were not collected from these animals, and aerobic cultures were not performed, those possibilities were not addressed in this study.

Further research regarding transmission probabilities and modes is needed. Larger sample sizes and balanced designs would allow more rigorous statistical analyses. However, establishing and maintaining such designs would be difficult, as tortoises' clinical, culture, and PCR status can vary over both the short and long term (e.g., see Chapter 4). Experiments entailing differences in pen design could be planned to estimate the probabilities of transmission by the various routes. For example, double layers of wire screening between pens, with sufficient distance between the layers to prohibit direct






74


contact, could be used to assess the probability of aerosol transmission. Pen designs in which water containers are shared, but direct contact and aerial transmission are precluded could generate data addressing the role of transmission via water. Tortoises' strength, determination, and digging abilities must all be considered when designing such experiments.

Seropositive, clinically healthy, culture and PCR negative individuals may be suitable for relocation, restocking, and repatriation programs, but clinically ill animals should not be used in such efforts. However, the proportion of tortoises that appear to have recovered from disease and eliminated the mycoplasma, while actually harboring the organism, is unknown. As evidenced in the preceding chapter, some of those tortoises may have latent infections that will recrudesce under stressful conditions.















CHAPTER 6
VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII



Introduction


Many mycoplasmas can be transferred vertically, that is, from mother to offspring in utero or in ovo (see Chapter 1). Before appropriate decisions can be made concerning the fate of desert or gopher tortoises infected with M. agassizii, particularly those from threatened or endangered populations, questions regarding the vertical transmissibility of M. agassizii need to be resolved. If M agassizii is not transmissible via eggs, the options for preserving genetic material are increased. I designed experiments to test the hypotheses that M. agassizii is transferred in ovo to gopher tortoise hatchlings and that levels of specific antibodies against M. agassizii in egg yolk and hatchling plasma are associated with those found in maternal plasma.


Methods


Adult tortoises were housed as detailed in Chapter 2, and selected as specified in Chapter 5. In order to obtain clutches of eggs, pairs were established in August, 1994, as detailed in Chapter 5. The sampling schedule was as detailed in Chapter 5.






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76


Egg Collection and Incubation

Egg development was monitored by radiography (6 mas, 62-76 kv) beginning in April of 1995 and 1996. When eggshells were judged to be calcified adequately, oviposition was induced by intravenous injection of arginine vasotocin (Sigma) at approximately 10-12 picograms/kg. Tortoises were monitored until clutch deposition was complete or for a minimum of 4 hr. Eggs were collected as they were laid and placed in a sanitized container (washed with 1:10 bleach solution and air-dried upside down) partially filled with sterilized vermiculite moistened with an equal weight of sterilized water. Female identification number and letter indicating order of deposition were written on each egg with a graphite pencil swabbed with 70% ethanol. After approximately half the clutch was laid, cloacal swabs for mycoplasma culture were taken, streaked onto SP4 agar, placed in SP4 broth, and incubated as described previously.

In 1995, eggs were incubated at 290C until hatching. In 1996, eggs were

incubated at 27, 29 or 3 1C. Approximately 1 wk before hatching each egg was cleansed of adhering vermiculite with clean gauze and placed into a sanitized plastic container. After hatching, resorption of yolk, and closing of the umbilicus, hatchlings were placed in containers with clean sand. Hatchlings were maintained separately from adults, at ambient temperature and light cycles, and fed natural foods supplemented with commercially available vegetables until release into outdoor pens, where they were fed natural foods. Culture and PCR Procedures

Eggs were taken at various times during incubation for mycoplasma culture and antibody detection. At pipping, chorioallantoic-amniotic fluid (CAF) was collected and






77


frozen for later analyses. For culture, 100 pl samples of yolk and albumin, and small pieces of membrane were added to 900 pl of SP4 broth. After 48 hr incubation at 300C, 500 pl of culture were removed for PCR analysis. All CAF samples obtained at pipping were treated in the same manner. In 1996, one or two pooled samples consisting of 100 pl of CAF from each of 3 - 6 eggs, mixed well, were made for each clutch and processed as described.


ELISA Procedures

Based on preliminary experiments using several methods of antibody extraction, the supernatant resulting from mixing 1 ml of yolk with 1 ml PBS-AZ provided the most efficient fraction for detection of antibodies (G. S. McLaughlin and I. M. Schumacher, unpub. data). Blood samples for ELISA were collected by cardiocentesis from most hatchlings at approximately 2-4 wk of age. In 1995, the goal was to obtain a blood sample from each hatchling. In 1996, sampling effort was reduced to a maximum of five samples per clutch due to the low coefficients of variation for the 1995 samples.

In 1995, hatchling ELISA samples were run as previously described except that three dilutions from the range 1:1 to 1:8 were run in duplicate, depending on the sample volume obtained. Sample dilutions were based on previous experiments with plasma from desert tortoise hatchlings (I. M. Schumacher, unpub. data). All hatchling or egg samples were run on the same plate as 1994 and 1995 samples from the corresponding female and the presumed sire, if possible. If samples were split between two plates, plates were run on the same day with the same reagents and positive and negative controls. In 1996, samples were run in triplicate at a dilution of 1:2, as that was the dilution used for analysis






78


of the 1995 samples. Additionally, that modification reduced the necessary blood volume from approximately 400 p to 150 pl, reducing stress to the hatchlings. Samples collected in 1995 and 1996 from the corresponding female were run on the same plate as samples from their eggs and/or hatchlings.


Results


Clutch Sizes, Fertility and Hatching Rates

Twenty-six clutches, 13 each year, were collected. Arginine vasotocin was used to induce oviposition of 20 clutches and the remaining six clutches (five in 1995 and one in 1996) were laid in the pens. In 1995, 13 tortoises developed 115 eggs (mean = 8.8), and 103 eggs were recovered (Table 6-1). Five females laid entire clutches in the pens, four of which were removed within 1 wk of laying. One female laid nine eggs in an inaccessible area of her pen. Two females that laid incomplete clutches on induction laid the remaining eggs (n = 3) in their pens. Twenty-five eggs were removed for culture, PCR, and ELISA samples. Of those, five were fertile and removed while viable, four had died, and five were infertile. I could not determine fertility on the remaining 11.

Overall clutch sizes (Table 6-1) were larger in 1996 (mean = 11.1) than in 1995 (mean = 8.8, 2-tail t-test = 2.78, 17 df P = 0.013), as were those of seronegative females (11.7 vs 9.8, 2-tail t-test = 2.45, 9 dfA P = 0.037). In 1996, clutch sizes of seronegative females (mean = 11.7) were larger than those of seropositive females (mean = 10.3, 2-tail t-test = 2.30, 11 dt P = 0.042). Thirteen females developed 144 eggs, 130 of which were recovered. One female laid 11 eggs that were incompletely calcified and broke during the






79


laying process. Five eggs were laid in the pen and six while in transport; a usable sample for culture and PCR was obtained from only one egg. Three females retained one egg each, the fates of which were unknown. One fertile egg was donated for the purpose of establishing embryonic tortoise cell lines, and 19 eggs were sampled. Of those, 13 eggs were infertile, four had died during incubation, and I could not determine fertility of two.

In 1995, 78 eggs pipped and 77 hatchlings were produced in the laboratory (Table 6-1), and nine hatchlings were recovered from the 12 eggs laid in the pens (1/1, 1/2, 7/9). The tortoise that pipped but did not hatch had an abnormal yolk sac; it was either malformed or had ruptured, and yolk was found throughout the coelom. The hatchling was in respiratory distress (cyanotic and mouth breathing), and was euthanatised. Another fetus appeared to be full term, but had spina bifida in the cervical region, and only one kidney.

In 1996, 111 eggs produced 113 viable hatchlings (Table 6-1). One egg produced one healthy hatchling and one that was anencephalic. Two other eggs, from a different clutch, each produced healthy twins. There were no differences in fertility or hatching rates between infected and uninfected females for either year individually or both years combined (X2 tests, 1 df all P > 0.75). Fertility and hatching rates did not differ significantly between years (X2 tests, 1 df, all P > 0.20). Culture and PCR Results

No cultures or PCR assays of cloacal samples, egg yolk, albumin, or membranes were positive for mycoplasma, and no hatchlings isolated from adults developed clinical signs (see Chapter 5 for evidence of horizontal transmission from adults to hatchlings).






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Table 6-1. Clutch data from gopher tortoise upper respiratory tract disease vertical transmission study, 1995 and 1996.
1995 1996
Serostatus negative positive total negative positive total
No. clutches 8 5 13 7 6 13 No. eggs 78 37 115 82 62 144
a a a,b b a
Mean 9.8 7.4 8.8 11.7 10.3 11.1
No. recovered 75 28 103 69 61 130 No. sampled 16 9 25 11 8 19
Fertility rates 94% 97% 95% 88% 92% 90%
No. hatchlings 61 26 87 52 61 113
Hatching rates 90% 93% 91% 86% 88% 87%
CAF samples 59 19 78 51 51 102 Blood samples 51 25 76 29 28 57
Clutch sizes differed significantly between 1995 and 1996 for seronegative tortoises (2-tail t-test = 2.45, P = 0.037), and for overall clutch size (2-tail t-test = 2.78, P =
0.013).
bClutch sizes differed significantly between seronegative and seropositive tortoises for
1996 (2-tail t-test = 2.30, P = 0.042).



ELISA Results

For ELISAs in 1995, 76 hatchlings and 34 yolks were sampled for a total of 96%

of all available eggs and hatchlings sampled. Total sampling effort per clutch ranged from

75-100%, with 50-100% of hatchlings sampled. In 1996, 57 hatchlings and 19 yolks were

sampled, for a total sampling effort of 58%, with sampling effort per clutch of 42-100%.

ELISA values in egg yolks and hatchlings were correlated with maternal ELISA values in

Autumn of the prior calendar year (r = 0.68, R2 = 0.461, P < 0.0002, n = 25; Figure 6-1),

indicating that specific antibodies were transferred via the egg yolks to the hatchlings.









1.00

y = 0.235x + 0. 1285
0.90 = 0.4611 P < 0.0002
0.80 R= 0.68 n =25
0.70 0.60

4 0.50

0.40

0.30


0.20

0.10 *

0.00
0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60 1.80 2.00
Maternal Antibody, OD @ 405 nm


Figure 6-1. Regression of specific anti-Mycoplasma agassizii antibody levels of gopher tortoise hatchlings on maternal antibody levels, as measured by ELISA. OD = optical density. 00






82

Discussion


Unlike various mycoplasmal infections of rodents and poultry (Simecka et al. 1992) and wild ducks (Goldberg et al. 1995), there was no evidence to support a hypothesis of vertical transmission of M. agassizii. Therefore, it should be possible to collect eggs from infected female tortoises, incubate the eggs, and release the hatchlings with no risk of spreading the disease.

Vitellogenesis begins in August or September and continues to December (Taylor 1982), providing an extensive period for deposition of antibodies. In spite of that, the antibody level in egg yolks and hatchling plasma was only 10 -20% of that in the maternal plasma. Thus, there is no evidence that female gopher tortoises are sequestering antibodies in the egg components. This is in contrast to birds, particularly chickens, which deposit large amounts of antibodies in their eggs in a very short time period. Antibody levels in chicken eggs can be several orders of magnitude greater than that in the maternal plasma.

Hatchling antibody levels decline slowly over the first year of life, as the maternal antibodies are broken down (I. M. Schumacher, unpub. data). This process is much slower in tortoises than in mammals and birds, where such passively acquired antibodies decline within weeks or a few months after birth or hatching. It is not known if antibodies against M. agassizii affect juvenile responses to infection with the organism. Although adult tortoises that have developed an antibody response against M. agassizii respond very quickly and adversely to subsequent exposure (Chapter 4), that response may be mediated more by cellular immune components (e.g., macrophages and heterophils) than by






83


humoral. Hatchling and juvenile tortoises with maternal antibodies may be protected from some effects of infection if the antibodies are neutralizing in nature, and if they can be mobilized appropriately against the mycoplasma. Research needs to be conducted to address these areas.















CHAPTER 7
ENVIRONMENTAL TRANSMISSION OF MYCOPLASMA AGASSIZII Introduction


Some mycoplasmas can remain viable for at least several days in the environment (Chandiramani et al. 1966), and for many years under refrigeration or freezing (Yoder and Hofstad 1964). One method of controlling mycoplasma infections in domestic stock is to depopulate a farm or facility, spray buildings and equipment with disinfectants, wait an appropriate amount of time, and then reintroduce uninfected stock (Anonymous 1989). In order to determine appropriate time frames for restocking tortoises, knowing the length of time Mycoplasma agassizii remains viable in the environment is critical. Unfortunately, direct viability testing is extremely difficult and impractical, if not impossible. Many soil bacteria and fungi in the tortoise burrow and surrounding environment grow very rapidly in culture and quickly overtake any mycoplasma colonies that might be present. However, the risk of environmental transmission of M. agassizii is an important parameter in the decision making process. In order to address this question, I designed an experiment to test the hypothesis that environmental transmission ofM. agassizii would occur in tortoises introduced to pens previously occupied by infected, clinically ill tortoises.








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Methods


As a result of previous and concurrent experiments, 13 seropositive tortoises

infected with M. agassizii and 15 seronegative tortoises with no clinical signs and negative culture and PCR results were available. The 13 positive tortoises were captured and reinoculated with approximately 108 colony forming units (CFU) ofM. agassizii to ensure active clinical disease and shedding of bacteria at the time scheduled for introducing healthy tortoises to the pens (see Chapter 4). Six weeks following inoculation, the 13 tortoises were captured, as were the 15 healthy tortoises. Nasal flush samples were collected from each infected tortoise to analyze by PCR.

Immediately after capture and removal of five infected tortoises from their pens, a healthy tortoise was put into the pen. Transfers were carried out within 15 min. Transfers were also carried out at 3 (n = 4), 7 (n = 3), and 10 (n = 3) days following removal of the infected tortoises. Because only 13 ill tortoise pens were available, one of the day 7 and one of the day 10 transfers consisted of pairs of tortoises. Eight weeks following transfer, the 15 tortoises were captured, assessed clinically, photographed, and blood and nasal flush samples were collected. Samples were handled as previously described.


Results


All infected tortoises were clinically ill at the time of capture, although PCR results for all animals were negative. Transferred tortoises explored their new surroundings and usually entered the burrows within 1 hr, and some entered within 15 min. At 8 wk posttransfer, no transfer tortoise was clinically ill, or had positive culture or PCR results. No tortoise showed an increase in ELISA values indicative of possible infection (Table 7.1).







86


Table 7.1. ELISA results from initially clinically healthy, culture and PCR negative tortoises transferred into pens previously occupied by clinically ill, culture or PCR positive tortoises. Ratios for equal ELISA values may differ due to plate--to--plate variation in the assay.
Tortoise Pre-transfer ELISA Post-transfer ELISA Transfer
ID Value Ratio Value Ratio Day
108 0.275 1.89 0.242 1.66 0 129 0.164 1.17 0.137 0.98 0 160 0.185 1.75 0.236 1.80 0 201 0.217 1.70 0.192 1.51 0 205 0.471 3.37 0.409 2.93 0

110 0.119 0.94 0.137 1.08 3 211 0.222 2.00 0.234 2.11 3 219 0.352 2.72 0.184 1.43 3 311 0.134 1.21 0.144 1.29 3

159 0.278 2.12 0.160 1.23 7 226 0.279 2.63 0.212 2.00 7 261 0.378 2.92 0.287 2.21 7

119 0.113 1.02 0.120 1.08 10 125 0.063 0.60 0.054 0.52 10 140 0.134 1.23 0.152 1.40 10



Discussion


Although the PCR results for the re-infected tortoises were negative at the time of

capture for transfer, the PCR is limited in its sensitivity. The lower limit of detection is

approximately 1000 CFU (D. R Brown, unpub. data), whereas the infectious dose of M.

agassizii may be 10 or fewer CFU (Brown et al. 1996b). Therefore, tortoises could have

been shedding infective doses of bacteria and still have negative PCR results.

In other studies, 100% of 28 naive tortoises seroconverted by 8 wk PI, with all

tortoises tested at 6 wk PI having significantly increased antibody levels (Brown, D. R.,

1996b; Schumacher, unpub. data; Chapter 4). No tortoises in this study had increased






87


antibody levels, and it is unlikely that insufficient time had passed for the tortoises to develop an immune response.

Environmental transmission of M. gallisepticum is of considerable concern to the poultry industry, necessitating disinfection of premises and equipment, and a 2-6 wk fallow period before introducing new stock (Anonymous 1989, M. B. Brown pers. comm.). The M. gallisepticum strain causing conjunctivitis in house finches also can survive in the environment and cause infections in individuals later housed in the same facility (Ley et at 1996, Luttrell et al. 1996).

Environmental transmission ofM. agassizii in the wild may not be of great

concern. However, equipment used for capturing, handling, holding, and transporting tortoises should be cleaned after each use by spraying or wiping with bleach, ethanol or other disinfectant solution. Care should be taken to dispose properly of all gloves or drapes that may have become contaminated. Clinically ill tortoises should not be housed in direct contact with other animals, nor should indirect contact be allowed. Infected tortoises should not be able to sneeze on other tortoises, nor should water or food dishes be shared between pens without disinfection. Because the sample sizes at each time point were very small, further research, with more rigorous attempts to isolate mycoplasmas and quantify shedding, needs to be conducted. For the purposes of relocation, restocking, or repatriation efforts, a short fallow period, perhaps 2 wk, is probably sufficient to prevent environmental transmission of M agassizii.















CHAPTER 8
CONSERVATION AND MANAGEMENT IMPLICATIONS OF UPPER RESPIRATORY TRACT DISEASE




The research reported in this dissertation has built on and extended the findings of research on upper respiratory tract disease in desert and gopher tortoises. Based on the knowledge that Mycoplasma agassizii causes clinical signs and lesions of URTD and elicits an antibody response detectable by an ELISA, experiments were designed to compare the pathological effects of the disease in gopher tortoises to those in desert tortoises, investigate secondary immune responses, and determine routes of transmission. The knowledge gained can be applied to conservation and management decisions, and new areas of research have been suggested.


Implications for Conservation and Management


When tortoises are impacted by development, mining, agriculture, or forestry

practices, decisions must be made regarding their disposition. Tortoises can be ignored, temporarily removed and confined to pens for later return to the site, permanently moved to a currently inhabited site, moved to a formerly occupied site, donated to research or educational facilities, or euthanatised. The choices open for a particular population of tortoises depend on the location, historical, current, and future site use, surrounding land



88






89


use patterns, importance of the population in maintaining genetic variability, and political and social factors. No one prescription will be ideal for all situations, and it is difficult, if not impossible, to develop a set of prescriptions that will cover all permutations of the above factors. However, guidelines for making decisions can be developed, and some are presented in this chapter.


Establishing Goals

The goals of the management action(s) must be established before decisions can be made regarding what data to collect, or how to design survey or monitoring programs. Management tactics designed to establish, create, or maintain a URTD-free population will differ from those intended to maintain the status quo relative to disease agents. If the genetic material represented by tortoises in defined or isolated populations (e.g., those in South Carolina, Mississippi, or Louisiana, and some in Alabama) is important for conservation purposes, aggressive efforts to protect and maintain the gene pool may be of primary concern.


Understanding URTD and Test Results

An understanding of test results is necessary for proper interpretation and

application. For field personnel, recognition of clinical signs of URTD is an important skill to gain. Differentiation of signs of URTD from other conditions is difficult even for experts, as the signs are nonspecific, but some conditions can be recognized fairly easily (e.g., foreign body in or trauma to the eye causing discharge, or nasal "discharge" due to






90


drinking or eating). Tortoise behavior interacts with clinical disease to affect transmission probabilities, so understanding behavior also is important in the decision making process.

A clinically healthy tortoise, with negative ELISA, culture, and PCR results is probably free from URTD. A positive ELISA, in the absence of clinical illness and positive culture or PCR results, indicates only that the tortoise has been exposed to M. agassizi. Because clinical signs and culture and PCR results can vary over time, we cannot predict if or when a seropositive tortoise will break with clinical disease and begin shedding bacteria. The more stress to which an animal is subject, such as human intrusion, handling or transport, drought or other extreme weather conditions, the more likely it is to have a disease recurrence. Repeated exposures to M. agassizii elicit more intense immunological responses by the tortoises, potentially leading to autoimmune responses that may contribute to the more severe lesions seen in longer-term infections, as well as possible liver pathology. Tortoises with rapid responses to the mycoplasma, with copious mucus production, may be more likely to transmit the agent to conspecifics, as their energy reserves have not been depleted by the disease process.

A clinically ill tortoise with positive culture or PCR results, regardless of ELISA

results, probably is capable of transmitting M. agassizii. The more active a tortoise is, and the greater its daily movements, the more likely it is to spread the bacteria through a colony and foment an outbreak of URTD. Behaviorally, male tortoises have larger home ranges and more intraspecific contacts, so are probably at greater risk of coming in contact with and spreading the pathogen.






91


A clinically healthy tortoise (i.e., one with no nasal or ocular discharge or other signs) may have positive culture and PCR results, with either positive or negative ELISA results. It may be in the early stages of infection or recrudescence. In the former case, the ELISA value may be negative, but should rise within 6 wk. In the latter case, the ELISA value may be quite high. Such tortoises may be capable of transmitting the mycoplasma under the appropriate conditions. Although direct contact (nose-to-nose) seems to be the most important route, transmission through water or on food cannot be ruled out. When tortoises drink, they often expel water through their noses for short distances, up to 50 cm, or sneeze forcefully after drinking, aerosolizing the contents of the nasal passages. If tortoises are in close contact with one another, spatially or temporally, such occurrences may allow transmission of M. agassizii. Tortoises with slight nasal discharge, virtually undetectable, can also aerosolize bacteria by sneezing.

Although long-term studies on the effect of URTD on survival of individuals and populations have not been conducted, the evidence from surveys of desert tortoise populations and from Sanibel Island indicate that the disease can have severe negative impacts on population viability. Declines of 25-50% over 1-3 yr, and of 30-90% over 10 yr, are catastrophic for species that take 10-20 yr to reach maturity and have recruitment rates of 1-2%. Without marked improvement in recruitment rates, affected populations are unlikely to recover within a reasonable time frame. Developing Questions and Conducting Surveys or Monitoring Programs

Third, questions related to management goals, and taking into account test interpretation and tortoise behavior, must be developed. Some of the most common




Full Text
16
pen-reared wild turkeys has resulted in decreased productivity (Rocke et al. 1988), there is
little evidence to indicate that infection with M. gallisepticum strains commonly occurring
in domestic fowl poses a threat to wild turkey populations (Luttrell et al. 1991), or that
wild turkey populations are important in the epizootiology ofM gallisepticum (Davidson
et al. 1988). Mycoplasma gallopavonis has been isolated from wild turkeys in Texas
(Rocke and Yuill 1987), South Carolina, Georgia (Luttrell et al. 1991), Colorado, New
Mexico, and Oklahoma (Fritz et al. 1992). Although lethal to experimentally infected
domestic turkey eggs (Rocke and Yuill 1987), the pathogenicity of M. gallopavonis to
wild turkeys has not been investigated. Mycoplasma synoviae and other, untyped,
Mycoplasma spp. were isolated from turkeys in Arizona (Fritz et al. 1992), but no
association with disease was found. As with M. gallisepticum, wild turkeys do not appear
to be important in the epizootiology of M. synoviae or M. me/eagridis (Davidson et al.
1988).
In 1993 an epizootic of polyarthritis occurred in juvenile farmed crocodiles
(Crocodylus niloticus) in Zimbabwe. The outbreak was characterized by high morbidity,
but low mortality. A mycoplasma was isolated, determined to be a previously
unrecognized species, and named M. crocodyli (Mohan et. al 1995). In 1995 a die-off of
captive adult American alligators (.Alligator mississippiensis) at a private facility in
Florida, with veiy high mortality, was associated with systemic infection with a different
species of mycoplasma, also previously unrecognized (Brown et al. 1996a). The name M.
lacerti has been proposed. In addition to M. agassizii, a second mycoplasma was found in
desert tortoises with evidence of URTD, including clinical signs, histologic lesions, and/or
positive ELISA tests (Brown et al. 1995). In a small pilot study, that organism was found


CHAPTER 6
VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII
Introduction
Many mycoplasmas can be transferred vertically, that is, from mother to offspring
in tero or in ovo (see Chapter 1). Before appropriate decisions can be made concerning
the fate of desert or gopher tortoises infected with M. agassizii, particularly those from
threatened or endangered populations, questions regarding the vertical transmissibility of
M. agassizii need to be resolved. IfM agassizii is not transmissible via eggs, the options
for preserving genetic material are increased. I designed experiments to test the
hypotheses that M. agassizii is transferred in ovo to gopher tortoise hatchlings and that
levels of specific antibodies against M. agassizii in egg yolk and hatchling plasma are
associated with those found in maternal plasma.
Methods
Adult tortoises were housed as detailed in Chapter 2, and selected as specified in
Chapter 5. In order to obtain clutches of eggs, pairs were established in August, 1994, as
detailed in Chapter 5. The sampling schedule was as detailed in Chapter 5.
75


67
The initially seronegative, clinically ill, culture and PCR positive male (183) had
seroconverted by October 1994. Based on his original samples from July 1994, and the
ELISA values of his October samples, he probably seroconverted within 2 wk of pairing,
and was considered as ELISA positive for the analyses (see below). In the ten
experimental pairs, eight initially asymptomatic members, two of which were originally
ELISA-positive and six of which were originally ELISA-negative, became moderately ill
with signs of URTD and had increased ELISA values, with the latter six seroconverting.
Two other tortoises (125 and 190) showed mild clinical signs that may have been
associated with mechanical irritation or environmental conditions, but did not seroconvert.
Of the four initially uninfected tortoises that showed no increase in antibody levels,
three did not have positive culture or PCR results. One (190) had only mild ocular signs,
and two (110 and 144) never showed clinical signs. The fourth (125), which consistently
showed mild signs and had a suspect PCR result in October 1994, was exposed to a
tortoise (287) that had positive samples only at necropsy. Based on observations, they
had only limited interactions. The serosuspect female (205), who was in contact with both
110 and 125, showed occasional, transient, mild clinical signs, but never had positive
cultures or PCR results, nor did her antibody levels increase. In fact at all sample times
after August 1994, she was classified as seronegative.
Of the 16 originally culture and PCR negative tortoises, 10 later had positive or
suspect culture or PCR results. Eight were exposed to clinically ill, culture or PCR
positive tortoises, seroconverted and had positive results, the ninth had positive culture
results only from necropsy samples, and the last was the individual (125) discussed above.
No tortoise became ill unless exposed to another ill tortoise.


101
Eisenberg, J. F. 1983. The gopher tortoise as a keystone species. Pp. 1-4 in Bryant, R.
J., and R. Franz (eds.). The gopher tortoise: A keystone species. Proceedings 4th
Annual Meeting Gopher Tortoise Council. Florida State Museum, Gainesville, FL.
Ellison, J. S., L. D. Olson, and M. F. Barile. 1992. Immunity and vaccine development.
Pp. 491-504 in: Maniloff, J., R. N. McElhaney, L. R. Finch, J. B. Baseman (eds.).
Mycoplasmas: molecular biology and pathogenesis. American Society for
Microbiology, Washington, D. C.
Foggie, A., G. E. Jones, and D. Buxton. 1976. The experimental infection of specific
pathogen free lambs with Mycoplasma ovipneumoniae. Research in Veterinary
Science 21: 28-35.
Forsyth, M. H., J. G. Tully, T. S. Gorton, L. Hinckley, S. Frasca, Jr., H. J. Van
Kruiningen, and S. J. Geary. 1996. Mycoplasma sturni sp. nov., from the conjunctiva
of a European starling (Stunia vulgaris). International Journal of Systematic
Bacteriology 46: 716-719.
Fox, E., J. Kuo, L. Tilling, and C. Ulrich. 1994. SigmaStat Statistical Software for
Windows Users Manual. Jandel Scientific, San Rafael, CA. 827 pp.
Fritz, B. A., C. B. Thomas, and T. M. Yuill. 1992. Serological and microbial survey of
Mycoplasma gallisepticum in wild turkeys (Meleagris gallopavo) from six western
states. Journal of Wildlife Diseases 28: 10-20.
Gamer, J. A., and J. L. Landers. 1981. Foods and habitat of the gopher tortoise in
southwestern Georgia. Proceedings of the Annual Conference of the South East
Association of Fish and Wildlife Agencies 35: 120-134.
Godley, J. S. 1989. A comparison of gopher tortoise populations relocated onto
reclaimed phosphate-mined sites in Florida. Pp. 43-58 in J. E. Diemer, D. R. Jackson,
J. L. Landers, J. N. Layne, and D. A. Wood (eds.). Gopher tortoise relocation
symposium proceedings. Florida Game and Fresh Water Fish Commission Nongame
Wildlife Program, Technical Report No. 5. Tallahassee, FL.
Goldberg, D. R., M. D. Samuel, C B. Thomas, P. Sharp, G. L. Krapu, J. R Robb, K. P.
Kenow, C. E. Korschgen, W. H. Chipley, M. J. Conroy, and S. H. Kleven. 1995.
The occurrence of mycoplasmas in selected wild North American waterfowl. Journal
of Wildlife Diseases 31: 364-371.
Hansen, K L. 1963. The burrow of the gopher tortoise. Journal of the Florida Academy
of Science 26: 353-360.
Hopkins, S. R., and H. W. Yoder, Jr. 1984. Increased incidence of air sacculitis in
broilers infected with Mycoplasma synoviae and chicken-passaged infectious
bronchitis vaccine vims. Avian Diseases 28: 386-396.


14
infections, and were infected with M. gallisepticum (Ley et al. 1996, Luttrell et al. 1996).
Since then, the organism has been associated with morbidity and mortality in other species
of passerines, including goldfinches (Carduelis tristis) (Nettles 1996), and was transmitted
to a blue jay (Cyannocitta cristata) in a rehabilitation facility (Ley et al. 1996).
Conjunctivitis and M gallisepticum also were found in house finches in Georgia (Luttrell
et al. 1996). The potential for this organism to spread over a large area due to the long
distances traveled by migratory birds, the mixed species flocks in which they congregate,
and local concentrations of many species around bird feeders is of great concern to many
ornithologists and ecologists, as well as poultry producers. There are indications that the
strain, while highly pathogenic to chickens, is not readily transmissible to poultry under
natural conditions (Nettles 1996). A new species of mycoplasma (M sturni) was isolated
from the conjunctiva of a European starling (Sturua vulgaris) with conjunctivitis found in
Connecticut during the epomitic (Forsyth et al. 1996). Although it was isolated in pure
culture, it was described as growing rapidly, whereas pathogenic mycoplasmas typically
grow slowly; therefore, it could have overgrown M. gallisepticum if that species had been
present. The pathogenicity of M. sturni needs to be investigated.
Three new species of mycoplasmas have recently been described from raptors in
Europe (Poveda et al. 1994). All were associated with respiratory diseases clinically and
histologically consistent with those caused by Mycoplasma spp., with lesions including
hyperplasia of mucous glands, lymphoid hyperplasia, and perivascular cuffing (Poveda et
al. 1990). The species were isolated from buzzards (Buteo buteo), saker falcons (Falco
cherrug), and griffon vultures (Gyps fulvus), and have been named, respectively, M.
buteonis, M. falconis, and M. gypis. Pathogenicity of these species, their distribution in


Hatchling antibody, OD @ 405 nm
Maternal Antibody, OD @ 405 nm
Figure 6-1. Regression of specific anti-Mycoplasma agassizii antibody levels of gopher tortoise hatchlings on maternal antibody levels,
as measured by ELISA. OD = optical density.


42
Table 3-1. Summary of ELISA, PCR, culture, and nasal histopathology results from
necropsied gopher tortoises from various locations in Florida, for determining infection
with Mycoplasma agassizii.
Group
Clinical
Signs
ELISA
PCR
Mycoplasma
Culture
Histopathology
Normal
( = 8)
0%
0%
0%
0%
0%
Natural Infection
(n- 15)
60%
85%
92%
80%
92%
Table 3-2. Summary of aerobic culture results from nasal cavity swabs of gopher tortoises
from Florida.
Group
Growth
Species
Percent
of isolates
Normal
very scant to
Staphylococcus spp.
45
(n = 5)
moderate
Gram-negative rods
25
-hemolytic Streptococcus sp.
10
non-hemolytic Streptococcus spp.
10
Corynebacterium spp.
10
Natural
very scant to
Corynebacterium spp.
33
Infection
heavy
Pasteurella testudinis
18
(=11)
Staphylococcus spp.
16
Gram-negative rods
15
Micrococcus sp.
11
Flavobacterium, Pseudomonas,
7
Lactobacillus, and others


TABLE OF CONTENTS
page
ACKNOWLEDGMENTS i
ABSTRACT viii
CHAPTERS
1 INTRODUCTION 1
Gopher Tortoise Natural History 1
Tortoises and Upper Respiratory Tract Disease 6
Mycoplasmal Respiratory Diseases in
Domestic Animals and Humans 10
Mycoplasmal Diseases of Wildlife 13
Chronic Manifestations of Mycoplasmal Infections 18
Project Overview and Specific Objectives 18
2 METHODS 21
Tortoises, Intake Procedures, Clinical Assessments
and Sampling Methods 21
Culture Procedures 22
PCR Procedure 22
ELISA Procedure 24
Study Group Assignment 25
Husbandry Procedures 26
Necropsy Procedures 27
Histopathology Procedures 28
Statistical Analyses 29
3 NATURALLY OCCURRING UPPER
RESPIRATORY TRACT DISEASE 30
Methods 30
Tortoises 30
Necropsy and Histology Procedures 31
Microbial Isolation 31
Electron Microscopy 32
Results 33
Normal Anatomy and Histology 33
Pathologic Findings 37
ELISA and PCR Results 40
Microbial Isolation Results 40
Discussion 40
v


I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Q. idL
Paul A. Klein
Professor of Pathology, Immunology and
Laboratory Medicine
This dissertation was submitted to the Graduate Faculty of the College of Agriculture
and to the Graduate School and was accepted as partial fulfillment of the requirements for
the degree of Doctor of Philosophy.
May 1997
guj\ X. <3*/
C2-
Dean, College of Agriculture
Dean, Graduate School


Absorbance @ 405 nm
Weeks Postinfection
Figure 4-3. ELISA results for three groups of tortoises, one control and two experimentally infected with Mycoplasma agassizii.
Tortoises in the naive group had no previous exposure to M agassizii, while those in the challenge group had evidence
of prior exposure. Line indicates ratio value (sample to negative control) of 3.0. Indicates significant differences
(P < 0.05) between challenge and naive groups. ** Indicates significant differences between naive and control groups.
*** Indicates the first significant differences from Time 0 value for that group. Bars represent standard errors.
4^


Along with increased physiological stress placed on tortoise populations from
human activities, there may be toxicological or immunological stress from chemicals
introduced into the environment. No research has been conducted into the effects of
herbicides, fungicides, insecticides, and fertilizers on gopher tortoise health, growth, or
reproduction. Investigations of disease in free-ranging populations of tortoises have
begun only recently.
Tortoises and Upper Respiratory Tract Disease
Although individual captive and wild gopher tortoises have been observed with
clinical signs of respiratory diseases for over 20 yr (E. R. Jacobson, unpublished data), the
first documentation of a larger-scale disease outbreak was in 1989, when an epizootic of
upper respiratory tract disease (URTD) was documented on Sanibel Island, Lee County,
Florida (G. S. McLaughlin and M. Elie, unpub. data). With the loss of 25-50% of
breeding-age adults in one population, recovery could take 50-150 yr (G. S. McLaughlin
unpub. data), barring further major losses and without substantial habitat improvement
leading to increased recruitment.
In the 1980s, large-scale population reductions (33-76% over 10 yr) of desert
tortoises (Gopherus agassizii) were documented at several sites in the western Mojave
Desert of California and at one site in the eastern Mojave (Com 1994, Berry in press).
Tortoises with clinical signs of URTD were observed among the remaining populations at
several sites (Knowles 1989; Berry 1990, in press). As a result of the declines, tortoises in
the Mojave Desert north and west of the Colorado River were declared threatened (U.S.
Fish and Wildlife Service 1990).


34
Figure 3-2. Diagrammatic representation of the interior of a gopher tortoise head
sectioned longitudinally, illustrating the relationship of the nasal cavity to the
external and internal nares. Approximately 3x life size. Drawing by L.
Mallory.


108
U. S. Fish and Wildlife Service. 1990. Endangered and threatened wildlife and plants;
determination of threatened status for the Mojave population of the desert tortoise.
Federal Register 55: 12178-12190.
Van Kuppeveld, F. J. M., J. T. M. Van Der Logt, A. F. Angulo, M. J. Van Zoest, W. G.
V. Quint, H. G. M. Niesters, J. M. D. Galama, and W. J. G. Melchers. 1992. Genus-
and species-specific identification of mycoplasmas by 16s r RNA amplification.
Applied Environmental Microbiology 58: 2602-2615.
Washburn, L. R., and E. Weaver. 1996. Protective effect of active and passive
immunization of rats against two surface antigens of mycoplasma arthritidis. IOM
Letters 4: 31.
Westhouse, R. A., E. R. Jacobson, R. K. Harris, K. R. Winter, and B. L. Homer. 1996.
Respiratory and pharyngo-esophageal iridovirus infection in a gopher tortoise
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Wilson, D. S. 1991. Estimates of survival for juvenile gopher tortoises, Gopherus
polyphemus. Journal of Herpetology 25: 376-379.
Witz, B. W., D. S. Wilson, and M. D. Palmer. 1991. Distribution of Gopherus
polyphemus and its vertebrate symbionts in three burrow categories. American
Midland Naturabst 126: 152-158.
Woldehiwet, Z., B. Mamache, and T. G. Rowan. 1990. Effects of age, environmental
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Tallahassee, FL. 14 pp.
Woodruff, R. E. 1982. Arthropods of gopher tortoise burrows. Pp. 25-48 in R. Franz
and R. J. Bryant (eds.). The gopher tortoise and its sandhill habitat. Proceedings of
the 3rd Annual Meeting Gopher Tortoise Council. Florida State Museum, Gainesville,
FL.
Wright, S. 1982. The distribution and population biology of the gopher tortoise
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Clemson, SC.


57
hosts proteins may sensitize the host and induce an autoimmune response, leading to
chronic manifestations of disease even if the primary etiologic agent is cleared.
Although actively produced antibodies apparently do not confer protection to adult
tortoises, it is unknown if passively transferred antibodies (see Chapter 6) provide
protection against URTD for hatchlings. Preliminary observations indicate that
seronegative hatchlings are at least as susceptible to infection as adults, and that the
disease progresses more rapidly, with high morbidity in the first 6 mo PI (see Chapter 5).
Desert tortoise hatchlings exposed to infected adults developed clinical signs of URTD
and suffered extremely high mortality rates (51 of 69, or 74%, of those with clinical signs
died) (Oftedal et al. 1996). Mycoplasma agassizii was isolated from necropsied
individuals, and severe inflammation was seen histologically. The immune status of the
hatchlings and parents was unknown prior to the development of the outbreak, so no
inferences can be drawn regarding the potential role of maternal antibodies in the course
of URTD in hatchlings. Because cellular components probably play a major role in the
pathogenesis of URTD and those are not transferred vertically in reptiles, the maternal
antibodies may provide some protection. It would be interesting to explore the question
of increased susceptibility to or severity of disease vs. protection from disease in
hatchlings that have received maternal antibodies against M. agassizii.
The immune response stimulated by previous exposure to M. agassizii did not
prevent reinfection or ameliorate signs of URTD after subsequent inoculation. The onset
of clinical signs was, in fact, more rapid and the initial severity was greater than in first
infections. Development of specific antibodies against M agassizii did not ensure
clearance of the organism by the tortoises, either on initial or subsequent exposure. Based


94
tortoises that may have cleared the bacteria would be at risk of developing severe disease
when exposed to a clinically ill tortoise, decreasing their survival probability.
If the goal is to establish, create, or maintain a URTD-ffee population, the
question remains of what to do with seropositive, clinically healthy, culture and PCR
negative individuals. If there is no secure place to hold the animals or a site where they
can be released, then the most acceptable choice may be euthanasia. Euthanasia may save
the animals from a long slow death, and prevents the chance of exposing negative animals.
Alternatively, intensive monitoring of the population, with special attention paid to
seropositive tortoises and samples collected several times per year, may allow rapid
detection and removal of ill animals. However, such careful monitoring is beyond the
capacity of most management agencies, and annual or biennial sampling may be all that
can be accomplished.
Researchers sometimes need seropositive animals for projects. If one can be
found, it is possible that animals could be donated. However, there are many issues to
consider with that approach, mostly for the researcher. Researchers usually cannot accept
animals that are made available suddenly, particularly for studies that require animals to be
kept in laboratory settings. Costs, objectives, confounding factors of length of disease,
genetics and site of origin, are all issues that must be addressed. It is expensive to keep
tortoises in captive research settings, and safeguards must be in place to prevent then-
escape or theft. Research can be a politically and socially sensitive subject due to
potentials for pain, illness, and death of animals, and nearly all animals are euthanatised at
the end of the studies.


83
humoral. Hatchling and juvenile tortoises with maternal antibodies may be protected from
some effects of infection if the antibodies are neutralizing in nature, and if they can be
mobilized appropriately against the mycoplasma. Research needs to be conducted to
address these areas.


Mean Clinical Sign Score
Weeks Postinfection
Figure 4-2. Total clinical sign scores for three groups of tortoises, one control and two experimentally infected with Mycoplasma agassizii.
Bars represent standard errors. ** Indicates a significant (P < 0.05) rise in clinical sign score and a significant difference
between groups. Indicates a significant rise in clinical sign score.


92
questions are: 1) What is the prevalence of antibodies against M. agassizii in the
population of interest? 2) What is the distribution of seropositive tortoises relative to
gender, age, habitat type, geography, or land use patterns? 3) Has the seroprevalence
changed over time? 4) Is clinical disease present on the sites? 5) Is M. agassizii
detectable from any tortoises? 6) Is there evidence, either from demographic profiles or
tortoise remains, of large die-offs, or gaps in recruitment?
Once the goals have been established and questions developed, surveys or
monitoring programs can be designed to collect the necessary samples and data. After
field work has been conducted, samples collected and analysed, and test results analysed,
the information can be used to develop management programs or research questions.
When trapping tortoises for studies or relocation, it is important to note any
clinical signs that are present. Simply being aware of the possibility that URTD exists on a
site may make field personnel more likely to detect clinical cases. While this does not
guarantee that every case of URTD will be caught, it could give an early warning of
potential problems, prevent sick tortoises from being put in holding pens with healthy
ones, or transmitting the agent on equipment. It is easy to spray equipment with a mild
bleach solution, or wipe down calipers and other equipment with alcohol soaked gauze. It
is more difficult, and expensive, to construct separate holding facilities for healthy and sick
tortoises, but alternatives are available. Ill tortoises can be kept in containers for a few
days, long enough for monitoring and until test results come back.


8 CONSERVATION AND MANAGEMENT IMPLICATIONS
OF UPPER RESPIRATORY TRACT DISEASE 88
Implications for Conservation and Management 88
Establishing Goals 89
Understanding URTD and Test Results 89
Developing Questions and Conducting Surveys
or Monitoring Programs 91
Weighing Management Options and
Formulating Management Plans 93
Summary of Conservation and Management Implications 95
Further Research 96
LITERATURE CITED 97
BIOGRAPHICAL SKETCH 110
vu


37
were seen. Golden brown granules were scattered throughout the kidney within renal
tubular epithelial cells of all tortoises. Mononuclear cells containing similar appearing
granules were within the renal interstitium and the interstitium of the testes.
Pathologic Findings
Gross examination of heads of diseased tortoises revealed minimal to large
amounts of exudate within the nasal cavity and nasal passageways. Histologically, of the
15 heads, one had no changes, two had mild changes, seven had moderate changes, and
five had severe inflammatory changes. In the tortoises with minimal changes, mild
mucosal hyperplasia and slightly increased lymphoid aggregates were seen in the nasal
passage and ventro-lateral depression. In those tortoises with moderate changes, the
olfactory epithelia were usually normal, with only focal or mild changes in the submucosa.
Changes generally were confined to the nasal passage and the ventral aspects of the nasal
cavities, and consisted of mucosal epithelial and lymphoid hyperplasia, with infiltration of
mononuclear cells and heterophils. In some tortoises with moderate inflammation, basal
cell proliferation and loss of cilia could be detected. In the tortoises with severe
inflammatory changes, there were lymphoid aggregates around submucosal glands, with
glandular epithelial hyperplasia. The normal mucosal architecture was replaced by
infiltrates of mononuclear cells and heterophils. The olfactory mucosa was replaced with
proliferating mucous epithelial cells (Figure 3-4). Proliferating basal cells projected into
the underlying lamina propria of some tortoises. Exudate, consisting of sloughed epithelial
cells and inflammatory cells, was found in the nasal cavity lumen.
Infected tortoises had larger lymphoid aggregates in the submucosa of the glottis;
several tortoises had focal areas of epithelial cell proliferation. Basal cell proliferation and


8
Hemoglobin concentration and phosphorous levels were lower in ill than in healthy
tortoises, and ill tortoises had higher sodium, blood urea nitrogen (BUN), creatinine,
serum glutamic oxalacetic transaminase (SGOT) activity, and total cholesterol levels.
Levels of liver and serum vitamins A and E did not differ between healthy and ill tortoises,
nor did liver values of selenium, copper, lead, or cadmium. However, iron and mercury
were higher in ill than in healthy tortoises. Healthy tortoises did not exhibit any
histological abnormalities in nasal mucosa or other tissues. Ill tortoises had less fat than
controls, atrophied thymuses, increased numbers of lymphocytes in the sinusoids of the
spleens, and increased amounts of iron granules in the hepatocytes. The most striking and
consistent changes were in the nasal passage and cavity tissues (Jacobson et al. 1991).
Grossly, passages and cavities of ill tortoises contained moderate to large amounts of
serous to purulent exudate. Histologically, the tissues exhibited loss of mucosal glands
and goblet cells, proliferation of epithelial cells, infiltrates of lymphocytes and histiocytes,
loss of cilia, dysplasia of the olfactory epithelia with infiltration of heterophils, basal cell
hyperplasia, occasional squamous metaplasia, and occasional erosion and ulceration of
mucosal epithelia. By electron microscopy, pleomorphic organisms resembling
Mycoplasma sp. were seen on cell surfaces and tightly adhered to cell membranes of ill
tortoises, but not healthy ones. Significantly more Pasteurella testudinis were isolated
from ill than healthy tortoises, and Bacillus sp. and Mycoplasma-like organisms were
isolated only from ill tortoises. By electron microscopy, the latter resembled those seen on
the nasal cavity tissues of ill tortoises (Jacobson et al. 1991).
The mycoplasma was determined to be a new species, provisionally named
Mycoplasma agassizii (Brown et al. 1994). An enzyme-linked immunosorbent assay


95
If the facilities and personnel are available, and the genetic material represented by
the sick or seropositive tortoises is important, then a captive breeding program may
provide the best solution. Clinically ill animals or those that test positive by culture or
PCR should be maintained separately from clinically healthy, culture and PCR negative
animals. Eggs can be collected from the females by induction of oviposition or at natural
laying, or nests can be dug up. Eggs should not be allowed to hatch and hatchlings
emerge from nests in pens inhabited by ill adults. Hatchlings can contract URTD from
adults, and they generally become very ill very quickly. Eggs can be artificially or
naturally incubated, and hatchlings released immediately or headstarted.
Summary of Conservation and Management Implications
In summary, the following points must be considered when making conservation
and management decisions relative to URTD in gopher tortoises:
1) Goals must be clearly established.
2) Personnel must have appropriate training to recognise URTD, collect necessary
samples, and interpret results.
3) Clear questions must be formulated.
4) Survey and monitoring programs must be developed and implemented, and
precautions taken to ensure detection and prevent spread of URTD.
5) Management options must be weighed, and plans formulated and implemented
that are consistent with established goals.
a) Habitat manipulations.
b) Relocate tortoises and maintain status quo relative to URTD.


25
sodium bicarbonate, pH 9.6, with 2 mM MgCl2, and added to wells at 100 pi/well. Plates
were incubated for 60 min in the dark, then read at 405 nm on a microplate reader (EAR
400 AT, SLT Labinstruments, Salzburg, Austria). The mean of two or three wells coated
with antigen and incubated with conjugate and substrate only was used as the blank. A
positive control, plasma from a naturally infected gopher tortoise from Sanibel Island, and
a negative control, plasma from an uninfected tortoise from Orange County, were included
on each plate.
Results from the ELISA were optical density (OD) readings from the microplate
reader. The OD readings reflected the intensity of the yellow color developed when all
components of the reaction (specific tortoise antibodies against M. agassizii, biotinylated
MAb HL673, AP-S, and /?NPP) were present. The readings are on a continuous scale, but
were interpreted categorically by calculating the ratio of the sample readings to the
negative control reading. Ratio values less than or equal to 2.0 were considered negative,
those greater than 2.0 and less than or equal to 3.0 were classified as suspect, and those
greater than 3.0 were classified as positive.
Study Group Assignment
Tortoises exhibiting one or more signs of disease, or with a positive culture or
PCR result, were designated as diseased. Animals were designated clinically healthy if
found free of signs of URTD, and with negative culture and PCR results. Tortoises were
assigned to study groups based on sex, weight, clinical assessments, culture, PCR and
ELISA results, and placed in the appropriate pens.


86
Table 7.1. ELISA results from initially clinically healthy, culture and PCR negative
tortoises transferred into pens previously occupied by clinically ill, culture or PCR positive
tortoises. Ratios for equal ELISA values may differ due to plate--to-plate variation in the
Tortoise
Pre-transfer ELISA
Post-transfer ELISA
Transfer
ID
Value
Ratio
Value
Ratio
Day
108
0.275
1.89
0.242
1.66
0
129
0.164
1.17
0.137
0.98
0
160
0.185
1.75
0.236
1.80
0
201
0.217
1.70
0.192
1.51
0
205
0.471
3.37
0.409
2.93
0
110
0.119
0.94
0.137
1.08
3
211
0.222
2.00
0.234
2.11
3
219
0.352
2.72
0.184
1.43
3
311
0.134
1.21
0.144
1.29
3
159
0.278
2.12
0.160
1.23
7
226
0.279
2.63
0.212
2.00
7
261
0.378
2.92
0.287
2.21
7
119
0.113
1.02
0.120
1.08
10
125
0.063
0.60
0.054
0.52
10
140
0.134
1.23
0.152
1.40
10
Discussion
Although the PCR results for the re-infected tortoises were negative at the time of
capture for transfer, the PCR is limited in its sensitivity. The lower limit of detection is
approximately 1000 CFU (D. R. Brown, unpub. data), whereas the infectious dose ofM
agassizii may be 10 or fewer CFU (Brown et al. 1996b). Therefore, tortoises could have
been shedding infective doses of bacteria and still have negative PCR results.
In other studies, 100% of 28 naive tortoises seroconverted by 8 wk PI, with all
tortoises tested at 6 wk PI having significantly increased antibody levels (Brown, D. R.,
1996b; Schumacher, unpub. data; Chapter 4). No tortoises in this study had increased


12
stressors (Whittlestone 1976). Lesions are alveolar, initially characterized by neutrophilic
infiltrates and later by lymphocytes and macrophages (Baskerville 1972).
Contagious bovine pleuropneumonia (CBPP), although not present in the United
States, is an economically important disease of cattle and water buffalo caused by M
mycoides subsp. mycoides (Howard and Gourlay 1978, Trichard et al. 1989). Morbidity
rates can reach 100% with up to 50% mortality, and recovered animals maintain a carrier
state. Neutrophilic exudate in the airways, serous exudate in the pleural cavity,
serofibrinous exudate in alveoli, and edema and necrosis of regional lymph nodes and
interlobular septa are seen at necropsy. Interseptal, peribronchial, and perivascular
lymphoid infiltration can be seen histologically (Hudson 1971). Calf pneumonia, with high
mortality among dairy calves, is caused primarily by viruses, but can be caused by either
M bovis or M. dispar (Stalheim 1983, Bryson 1985). Infection with the former is
characterized by peribronchiolar and alveolar monocyte infiltration, while the latter causes
interstitial pneumonia with monocyte infiltration of alveolar walls, but not peribronchiolar
infiltration. Mycoplasma dispar also causes superficial and asymptomatic infection of
respiratory mucosa (Woldehiwet et al. 1990).
Lambs are susceptible to infection with M. ovipneumoniae, which is transmitted to
neonates from the ewes. Pneumonia, characterized by coughing, sneezing, nasal
discharge, fatigue, and poor weight gain, can develop as colostral antibodies wane.
Lesions are characterized by alveolar proliferation, nodular lymphoid hyperplasia, and
peribronchial and perivascular lymphoid infiltration (Carmichael et al. 1972, Foggie et al.
1976). Severity can be exasperated by infection with Pasteur ella haemolytica biotype A
(Jones et al. 1982).


4
than C.I.T.E.S., those take precedence. Legal protection is extended to the species in all
states within the range, although the levels of protection vary. The populations west of
the Tombigbee and Mobile Rivers in Alabama, Mississippi and Louisiana are on the
federal threatened species list, which prohibits the taking, exportation, or interstate
movement of individuals originating from that region without permit, and the possession,
transportation, purchase, and/or sale of illegally obtained specimens. Permits issued by the
U. S. Department of Interior can be obtained for scientific research on wild populations,
and possession of limited numbers of individuals for exhibition, education, and/or research.
State permits are required also. The gopher tortoise is fisted in Alabama as a protected
nongame species, in Mississippi as endangered, and in Louisiana as threatened. Louisiana
regulations also prohibit the use of gasoline, chemicals, or volatile substances to flush
reptiles from burrows or other hiding places. Georgia fists the species as threatened and
issues scientific collection permits only to qualified institutions and individuals for
educational or research purposes. Although the law states that burrows may not be
disturbed nor destroyed, nor may explosives, chemicals or smoke be introduced into them
to drive out wildlife, the code explicitly exempts poisonous snakes. Because venomous
snakes, particularly rattlesnakes, use gopher tortoise burrows, tortoises can be adversely
impacted by such activities. South Carolina was the last state to extend legal protection to
gopher tortoises (Mann 1990, Levell 1995), and now fists the species as endangered, with
permits required for any activities involving tortoises. In Florida, where the gopher
tortoise is fisted as a species of special concern (Wood 1996), regulations prohibit taking
and disturbing of tortoises and their habitats, although exceptions are granted regularly for
development, agriculture, and mining operations. In the past, human consumption of


CHAPTER 4
EFFECTS OF REPEATED EXPOSURE ON SEROPOSITIVE ADULTS
Introduction
In order to properly evaluate the results of the diagnostic tests and incorporate
those findings into management and conservation plans, epidemiological questions,
including the response of seropositive, asymptomatic tortoises to subsequent exposure to
the agent, must be addressed. Although vaccines have been developed for some
mycoplasmal diseases (Ellison et al. 1992, Lai et al. 1996, Markham et al. 1996), they are
usually not fully protective (e.g., Djordjevic et al. 1996, Kleven et al. 1996, Mohan et al.
1996, Washburn and Weaver 1996), and many natural infections by mycoplasmas do not
engender a protective host immune response. The immune response is actually essential to
the development of lesions, and infected animals are susceptible to repeated infection
(Simecka et al. 1992). Due to the immunopathology, the disease may be more severe on
subsequent exposure than on initial infection. I designed an experiment to test the
hypothesis that tortoises that have produced antibodies against M. agassizii are protected
against reinfection with the organism and subsequent development of URTD.
46


23
supernatant aspirated. Three to four microliters of 20 pg/ml proteinase K (Sigma, St.
Louis, MO) in 20 pi lysis buffer (100 mM tris pH 7.5, 6.5 mM DTT, 0.05% Tween 20)
were added to the pellets, which were resuspended, and the samples were incubated at
37C for 8-16 hours. After denaturing the proteinase K at 97C for 15 min, 5 pi of each
sample were removed and added to 45 pi of reaction solution containing two primers for
the 16S rRNA gene at 1 pM each, deoxynucleoside triphosphates at 200 pM, 2.0 mM
MgCl, and 2.5 units of Taq polymerase (Promega, Madison, WI). 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', Brown et al.
1995) and a Mycoplasma genus-specific region [anti-sense strand nt 1055 to 1031,
5'-TGCACCATCTGTCACTCTGTTAACCTC-3', Van Kuppeveld 1992], Samples were
subjected to 50 cycles of template denaturation for 45 sec at 94C, primer annealing for 1
min at 55C, and polymerization for 45 sec at 72C, followed by 10 min at 72C. Positive
samples yielded 576 base pair (bp) products that were visualized by combining 15 pi of
product with 2 pi bromphenol blue in 50% glycerol solution and electrophoresing on
ethidium bromide-stained 1.5% agarose gels in tris-borate-EDTA buffer. Positive control
samples using 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, Hind III digest of x phage DNA, was included on each gel.
In order to confirm that the isolates obtained from naturally and experimentally
infected tortoises were M. agassizii, an additional procedure, restriction fragment length
polymorphism (RFLP) analysis, was conducted on at least one isolate from each tortoise.
Twenty microliter samples of products from the above amplification procedure were


term infections. Although clinically healthy, ELISA-positive, culture and PCR-negative
tortoises may have eliminated the bacteria, when five such animals were examined at
necropsy, three were found to harbor M. agassizii in the nasal cavities. When seropositive
tortoises were challenged with M. agassizii, a more rapid and more severe clinical
response resulted than on initial exposure, and plasma antibody levels began rising more
quickly. When uninfected animals were housed with infected individuals, horizontal
transmission occurred, probably via direct contact, but possibly via food, water, or
fomites. Transmission was more likely to occur from a tortoise that was clinically ill and
culture or PCR-positive. There was no discernible transmission when tortoises inhabited
pens or entered burrows previously occupied by ill tortoises. There was no demonstrable
vertical transmission, although there was transfer of maternal antibodies via egg yolk. The
level of antibodies in egg yolk or hatchling plasma was approximately 10-20% of that in
maternal plasma. Movement of tortoises during relocation, repatriation, or restocking
efforts potentially could transport M. agassizii to previously uninfected sites. Because of
the uncertainty involved in determining latent infections, ELISA-positive animals should
not be moved to locations with no seropositive individuals. However, they can be used in
captive breeding efforts and their offspring released into the wild. Clinically ill animals
should not be relocated to new sites, and should not be housed with clinically healthy
animals in temporary holding situations, such as on-site relocations, or in captive breeding
programs.
IX


35
Figure 3-3. Photomicrographs of normal gopher tortoise upper respiratory tract tissues.
Hematoxylin and eosin staining, 320x. a) Photomicrograph of the ventro
lateral depression demonstrating the ciliated mucous epithelium; b)
Photomicrograph of the dorsal nasal cavity demonstrating the multilayered
olfactory epithelium. Photographs by E. R. Jacobson.


CHAPTER 5
HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII
Introduction
In order to develop management and conservation plans that incorporate the
potential of disease to affect populations, epidemiologic questions must be addressed. Of
particular concern are the probability of transmission of the disease organism from one
tortoise to another (horizontal transmission), the rate of spread within populations, and the
potential for spread to nearby populations. The probability of transmission may be related
to a tortoises clinical and culture or PCR status. Infected tortoises may be culture and/or
PCR positive without showing clinical signs. Conversely, it is sometimes difficult to
detect bacteria in clinically ill tortoises. I designed an experiment to test the hypothesis
that horizontal transmission of M. agassizii will occur only from those individuals that are
clinically ill, and PCR and/or culture positive.
Methods
Acquisition of tortoises, intake and husbandry procedures, clinical assessments,
sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed
in Chapter 2.
59


79
laying process. Five eggs were laid in the pen and six while in transport; a usable sample
for culture and PCR was obtained from only one egg. Three females retained one egg
each, the fates of which were unknown. One fertile egg was donated for the purpose of
establishing embryonic tortoise cell lines, and 19 eggs were sampled. Of those, 13 eggs
were infertile, four had died during incubation, and I could not determine fertility of two.
In 1995, 78 eggs pipped and 77 hatchlings were produced in the laboratory (Table
6-1), and nine hatchlings were recovered from the 12 eggs laid in the pens (1/1, 1/2, 7/9).
The tortoise that pipped but did not hatch had an abnormal yolk sac; it was either
malformed or had ruptured, and yolk was found throughout the coelom. The hatchling
was in respiratory distress (cyanotic and mouth breathing), and was euthanatised. Another
fetus appeared to be full term, but had spina bifida in the cervical region, and only one
kidney.
In 1996, 111 eggs produced 113 viable hatchlings (Table 6-1). One egg produced
one healthy hatchling and one that was anencephalic. Two other eggs, from a different
clutch, each produced healthy twins. There were no differences in fertility or hatching
rates between infected and uninfected females for either year individually or both years
combined (yC tests, 1 df, all P > 0.75). Fertility and hatching rates did not differ
significantly between years (y2 tests, 1 df, all P > 0.20).
Culture and PCR Results
No cultures or PCR assays of cloacal samples, egg yolk, albumin, or membranes
were positive for mycoplasma, and no hatchlings isolated from adults developed clinical
signs (see Chapter 5 for evidence of horizontal transmission from adults to hatchlings).


26
Husbandry Procedures
Tortoises were housed singly or in pairs in outdoor pens at the UF Animal
Resource Farm (ARF). There were four groups of 10 pens in a larger, chain link-fenced
enclosure (Fig. 2-1). Pens were approximately 21 m2, constructed of a wooden frame
with sheet metal extending vertically approximately 0.7 m above and below ground,
partially covered by shade cloth, and provided with an artificial burrow, a water dish, and
a cement feeding stone. Pens were observed daily by ARF staff, and watered daily in the
summer and as needed in the winter. The tortoises were fed a salad of mixed vegetables
three times per wk, and fruit was provided on an occasional basis. In addition, I observed
the pens and tortoises from three to seven days each week, with some days including
multiple observations. Because of individual behavior patterns, not every tortoise was
observed at each time point. The amount of food eaten was recorded for each pen the day
following feeding, and the stones were cleaned. Remaining food was collected and bagged
separately for each section, using brushes and dust pans assigned to that section. Stones
were then sprayed with a dilute (1:30) bleach solution, allowed to soak for a few minutes,
and hosed off. Water dishes were rinsed and filled daily, and bleached and scrubbed as
necessary. 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 a different tortoise. Used gloves were placed in plastic garbage bags and
disposed of properly. Before entering the first pen on a given day, personnel sprayed their
footwear with a dilute bleach solution. Footwear was sprayed with bleach before leaving


24
incubated with 10-20 units of the endonuclease Agel (New England Biolabs, Inc.,
Beverly, MA), which cuts the M agassizii amplification product at nt 613, and 5 pi of
reaction buffer, at 25C for one hour, and the products electrophoresed as above. The
procedure resulted in products of 434 and 142 bp from M. agassizii-positive samples, and
no change in non-M agassizii- samp les.
ELISA Procedure
An aliquot of plasma from each sample was used for determinations of levels of
antibodies specific for M agassizii (Schumacher et al. 1993). Ninety-six-well microtiter
plates (Maxisorp F96, Nunc, Kamstrup, Denmark) were coated with 50 pi of a whole-cell
lysate of M. agassizii strain 723 at 10pg/ml in phosphate buffered saline with 0.02% azide
(PBS-AZ). Plates were incubated overnight at 4C, 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 4C with 250 pl/well PBST containing 5%
non-fat dry milk (PBS-TM). Following washing, 50 pi of plasma diluted appropriately for
the specific study with PBS-TM were added to individual wells in duplicate or triplicate,
and the plates were incubated at room temperature for 60 min. The plates were washed,
50 pl/well of a biotinylated monoclonal antibody (MAb HL673) against the light chain of
desert tortoise immunoglobulins IgY and IgM at 1 pg/ml in PBS-TM was added, and
plates were incubated for 60 min. Following washing, a conjugate of alkaline phosphatase
and streptavidin (AP-S; Zymed Laboratories, Inc., San Francisco, CA) at 1:2000 in PBS-
AZ was added at 50 pl/well, plates were incubated for 60 min, and washed. Substrate, p-
nitrophenyl phosphate disodium (pNPP; Sigma), was prepared at 1 mg/ml in 0.01 M


106
Nunoya, T., M. Tajima, T. Yagihashi, and S. Sannai. 1987. Evaluation of respiratory
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Oboegbulem, S. I. 1981. Enzootic pneumonia of pigs: a review. Bulletin of Animal
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Oftedal, O. T., T. E. Christopher, and M. E. Allen. Upper respiratory tract disease in
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19
experimental infections to determine conclusively the etiologic agent and dose-related
effects, 3) to refine the PCR test and delineate further the taxonomy of the tortoise
mycoplasmas, 4) to characterize the immune responses of gopher tortoises to URTD
pathogens, and 5) to compare the efficacy of culture, PCR and serology in detecting new
infections.
As the original experiments were conducted and concern over the continued well
being of tortoise populations throughout the range increased, additional questions were
raised. Those were concerned primarily with management and conservation of gopher
tortoises, particularly with respect to relocation, repatriation, and restocking efforts.
Several questions were raised repeatedly in meetings with state and federal agency
personnel, particularly regarding possible immunity to the organism and factors related to
transmission. In order to address those questions additional objectives were established,
as delineated in specific objectives 2-7 below, and discussed in Chapters 3 through 8.
Chapter 2 will detail the methods common to fulfilling the objectives.
This dissertation addresses the following specific objectives:
1) To describe the clinical presentation and lesions of natural infections by M. agassizii in
gopher tortoises and compare the lesions to those found in desert tortoises (Chapter 3).
2) To test the hypothesis that tortoises that have produced antibodies against M. agassizii
are protected against reinfection with the organism and subsequent development of
URTD (Chapter 4).
3) To assess the probability of horizontal transmission of M. agassizii between adult
tortoises and to test the hypothesis that transmission is more likely to occur from an


BIOGRAPHICAL SKETCH
GRACE SHERYL McLAUGHLIN
I spent most of my life on the West Coast of North America, and attended
Humboldt State University, Areata, California, receiving a Bachelor of Arts degree in
Zoology, in spite of spending all my time in the Wildlife Department. I then ran an organic
farm for several years before moving to Ames, Iowa, to attend Iowa State University. I
studied gopher tortoises on Sanibel Island, Florida, en route to a Master of Science degree
in Animal Ecology. Before defending my thesis in November 1990,1 began my doctoral
studies at University of Florida. I originally studied parasites of bobcats and Florida
panthers, particularly their hookworms. However, I accepted a job with the gopher
tortoise upper respiratory disease project, and switched dissertation research projects. I
hope to move west and/or north soon after completion of my work on the project.
110


45
northeast Florida, and seropositive tortoises at a site in Mississippi (D. M. Epperson, pers.
comm.). Although many Florida and Georgia tortoise populations are fairly large, those in
Mississippi are more restricted and are on the federal endangered species list. Outbreaks
of URTD in populations with limited recruitment and no nearby source populations could
contribute to severe declines in numbers and, possibly, local extinctions. Further studies,
on a range-wide basis, need to be conducted to determine the distribution and potential
impacts of URTD on gopher tortoise populations.
In addition, further research is necessary regarding the pathogenesis of URTD.
While findings of liver, kidney, and intestinal tract lesions are interesting, they do not
elucidate the mechanisms by which the changes are caused. Changes have been found in
hormonal profiles of some infected desert tortoises (Rostal et al. 1996), which could lead
to altered foraging and reproductive behavior, as well as decreased reproductive potential.
If foraging behavior is affected, food and water intake might be reduced, which could
affect liver and kidney functions. Both direct and indirect pathogenic mechanisms need to
be studied in order to better predict the effects of URTD on tortoises.


17
to cause disease and seroconversion in gopher tortoises previously unexposed to either
mycoplasma (D. R. Brown, et al., unpub data).
Mycoplasmas have been isolated from bighorn sheep (Ovis canadensis) with
pneumonia associated with P. haemolytica, and there is concern that these may have a role
in the disease process (D. Hunter, pers. comm.). A mycoplasma isolated from dead and
live captive bighorn sheep with pneumonia was typed asM arginini (Woolf et al. 1970).
Apparently, no further work has been done regarding the pathogenicity of that strain to
bighorn. With continued exposure of bighorn to domestic livestock, Mycoplasma spp.
could interfere with recovery efforts. An outbreak of pneumonia associated with M.
ovipneumoniae infection occurred in captive Dalis sheep (Ovis dalli dalli) following
indirect exposure to domestic sheep (Black et al. 1988). Although Dalis sheep are
unlikely to be exposed to domestic livestock in their native habitat, it is clear that captive
herds or those in transit must be protected from exposure to pathogens.
Behymer et al. (1989) detected antibodies to Mycoplasma spp. in mule deer
(Odocoileus hemiotms) in California, but there was no association with disease, and
isolations were not attempted. One-humped camels (Camelus dromedarius) and African
buffalo (Syncerm caffer) on a game farm in Kenya had antibodies against Mycoplasma
strain F38, and camels were seropositive for M. mycoides mycoides, but no isolations
were made and no disease was seen (Paling et al. 1988). Kirchoff et al. (1996) found, in a
survey of captive arthritic elephants (Elephas maximus and Loxodonta africana), that
about 60% of the females harbored a new mycoplasma in the urogenital tract, although
none was recovered from males. It is not known if the mycoplasma, named M. elephantis,
causes arthritis.


80
Table 6-1. Clutch data from gopher tortoise upper respiratory tract disease vertical
transmission study, 1995 and 1996,
1995
1996
Serostatus
negative
positive
total
negative
positive
total
No. clutches
8
5
13
7
6
13
No. eggs
78
37
115
a
82
a,b
62
b
144
a
Mean
9.8
7.4
8.8
11.7
10.3
ll.l
No. recovered
75
28
103
69
61
130
No. sampled
16
9
25
11
8
19
Fertility rates
94%
97%
95%
88%
92%
90%
No. hatchlings
61
26
87
52
61
113
Hatching rates
90%
93%
91%
86%
88%
87%
CAF samples
59
19
78
51
51
102
Blood samples
51
25
76
29
28
57
aClutch sizes differed significantly between 1995 and 1996 for seronegative tortoises
(2-tail t-test = 2.45, P = 0.037), and for overall clutch size (2-tail t-test = 2.78, P =
0.013).
bClutch sizes differed significantly between seronegative and seropositive tortoises for
1996 (2-tail t-test = 2.30, P = 0.042).
ELISA Results
For ELISAs in 1995, 76 hatchlings and 34 yolks were sampled for a total of 96%
of all available eggs and hatchlings sampled. Total sampling effort per clutch ranged from
75-100%, with 50-100% of hatchlings sampled. In 1996, 57 hatchlings and 19 yolks were
sampled, for a total sampling effort of 58%, with sampling effort per clutch of42-100%.
ELISA values in egg yolks and hatchlings were correlated with maternal ELISA values in
Autumn of the prior calendar year (r = 0.68, R2 = 0.461, P < 0.0002, n = 25; Figure 6-1),
indicating that specific antibodies were transferred via the egg yolks to the hatchlings.


UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES,
GOPHERUS POLYPHEMUS:
PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION,
AND IMPLICATIONS FOR CONSERVATION AND MANAGEMENT
BY
GRACE SHERYL MCLAUGHLIN
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1997

DEDICATION
This dissertation, the years that went into it, and the drive for knowledge
that underpinned it, are a result of, and dedicated to, my parents:
Grace Slater Gibson Billie McLaughlin
1928-1978
and
Robert Douglas McLaughlin
1925-1994
Thank you.

ACKNOWLEDGMENTS
I thank Dr. Don Forrester for accepting me as a graduate student, and for his
support and guidance through the years. Drs. Kathy Ewel and Paul Gibbs assisted my
growth scientifically and professionally in the first half of my program. Dr. Ewels support
was instrumental in obtaining my fellowship, and Dr. Gibbs was responsible for a trip to
Australia. Drs. Mel Sunquist and Wiley Kitchens weathered the changes in my project
with grace and humor, and Dr. Kitchens was especially helpful in teaching me to argue my
positions and not back down when I knew I was right. It took several years for Dr. Mary
Brown to get me into her lab, and her support in presenting me to her colleagues is
appreciated. Dr. Elliott Jacobson has done his best to teach me clinical pathology and
histopathology and has been very supportive of my contributions to the overall project.
Dr. Paul Klein has given me some valuable insights into the critical thinking process.
Without my co-workers Drs. Dan Brown and Isabella Schumacher, Sylvia Tucker,
Barbara Crenshaw, and Cathie McKenna, and technicians Alyssa Whitemarsh, Michael
Lao, and Dave Bunger, this research would have been impossible. I benefited from Dans,
Isas and Barbs teaching abilities, and their willingness to discuss theory, practical
applications, philosophical underpinnings and differences of opinion. Mr. Clement
Lindsey and his staff cared for my research animals.
I thank Dr. Tim Gross for all his help with eggs and hatchlings, and John Wiebe
and Carla Weiser for their care. Drs. Dale Jackson and Michael Ewert provided advice.
Drs. Bruce Homer and Claus Buergelt have given me more strength and support than they
m

knew and taught me much. Many of the VMTH personnel, especially Neil, Danielle, Tom,
Dana, and An have assisted and taught me, and cheered me up when I needed it.
Ms. Joan Berish has provided an incredible amount of advice, support, and
friendship throughout the eight years I have been studying gopher tortoises, and I only
hope to reciprocate. Garry Fosters assistance through my early years at UF were
invaluable, as were the support of my fellow graduate students in Dr. Forresters lab,
Marisol Sepulveda and Don Coyner. My brother Mark, my sister Megan, and friends
Vicky, Bill, Sharon, Kay, Pierre, Andrea, and Tania helped me get here and stick it out,
and deserve thanks for their support. There have been several other people who have
helped me at various times and in different ways, especially those of the gopher tortoise
sodality, and my thanks go to them also. I also thank the members and friends of the
Unitarian Universalist Fellowship of Gainesville for being my family and caring for me.
I was supported by a Presidential Research Fellowship and a Gatorade Grant from
the University of Florida, and funds from The Walt Disney World Company.
IV

TABLE OF CONTENTS
page
ACKNOWLEDGMENTS i
ABSTRACT viii
CHAPTERS
1 INTRODUCTION 1
Gopher Tortoise Natural History 1
Tortoises and Upper Respiratory Tract Disease 6
Mycoplasmal Respiratory Diseases in
Domestic Animals and Humans 10
Mycoplasmal Diseases of Wildlife 13
Chronic Manifestations of Mycoplasmal Infections 18
Project Overview and Specific Objectives 18
2 METHODS 21
Tortoises, Intake Procedures, Clinical Assessments
and Sampling Methods 21
Culture Procedures 22
PCR Procedure 22
ELISA Procedure 24
Study Group Assignment 25
Husbandry Procedures 26
Necropsy Procedures 27
Histopathology Procedures 28
Statistical Analyses 29
3 NATURALLY OCCURRING UPPER
RESPIRATORY TRACT DISEASE 30
Methods 30
Tortoises 30
Necropsy and Histology Procedures 31
Microbial Isolation 31
Electron Microscopy 32
Results 33
Normal Anatomy and Histology 33
Pathologic Findings 37
ELISA and PCR Results 40
Microbial Isolation Results 40
Discussion 40
v

4 EFFECTS OF REPEATED EXPOSURE ON
SEROPOSITIVE ADULTS 46
Introduction 46
Methods 47
Statistical Analyses 47
Experimental Design 47
Results 49
Clinical Signs 49
Culture and PCR Results 53
ELISA Results 53
Histology Results 55
Discussion 55
5 HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII . 59
Introduction 59
Methods 59
Experimental Design 60
Results 64
General Observations 64
Evidence of Transmission of Mycoplasma agassizii 66
Transmission Probabilities 68
Discussion 71
6 VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII 75
Introduction 75
Methods 75
Egg Collection and Incubation 76
Culture and PCR Procedures 76
ELISA Procedures 77
Results 78
Clutch Sizes, Fertibty and Hatching Rates 78
Culture and PCR Results 79
ELISA Results 80
Discussion 82
7 ENVIRONMENTAL TRANSMISSION OF
MYCOPLASMA AGASSIZII 84
Introduction 84
Methods 84
Results 85
Discussion 86
vi

8 CONSERVATION AND MANAGEMENT IMPLICATIONS
OF UPPER RESPIRATORY TRACT DISEASE 88
Implications for Conservation and Management 88
Establishing Goals 89
Understanding URTD and Test Results 89
Developing Questions and Conducting Surveys
or Monitoring Programs 91
Weighing Management Options and
Formulating Management Plans 93
Summary of Conservation and Management Implications 95
Further Research 96
LITERATURE CITED 97
BIOGRAPHICAL SKETCH 110
vu

Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the Requirements
for the Degree of Doctor of Philosophy
UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES, GOPHERUS
POLYPHEMUS: PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION, AND
IMPLICATIONS FOR CONSERVATION AND MANAGEMENT
By
GRACE SHERYL MCLAUGHLIN
May 1997
Chair: Donald J. Forrester, Ph. D.
Cochair: Mary B. Brown, Ph. D.
Major Department: Wildlife Ecology and Conservation
Upper respiratory tract disease (URTD) of tortoises is caused by the molhcute
Mycoplasma agassizii, and is characterised by nasal and ocular discharge, palpebral
edema, and conjunctivitis. Hyperplasia and dysplasia of the nasal passage and cavity
epithelia and inflammatory infiltrates are seen histologically. In order to provide data for
management decisions and to better understand URTD, I studied uninfected, naturally
infected, and experimentally infected tortoises. The pathological and immune responses of
tortoises to and transmission of M. agassizii were investigated using clinical and
histological observations, culture and polymerase chain reaction (PCR) tests, and an
enzyme-linked immunosorbent assay (ELISA). Infection with M. agassizii caused mild to
severe damage to mucosal and olfactory nasal epithelia, with increased damage in longer-

term infections. Although clinically healthy, ELISA-positive, culture and PCR-negative
tortoises may have eliminated the bacteria, when five such animals were examined at
necropsy, three were found to harbor M. agassizii in the nasal cavities. When seropositive
tortoises were challenged with M. agassizii, a more rapid and more severe clinical
response resulted than on initial exposure, and plasma antibody levels began rising more
quickly. When uninfected animals were housed with infected individuals, horizontal
transmission occurred, probably via direct contact, but possibly via food, water, or
fomites. Transmission was more likely to occur from a tortoise that was clinically ill and
culture or PCR-positive. There was no discernible transmission when tortoises inhabited
pens or entered burrows previously occupied by ill tortoises. There was no demonstrable
vertical transmission, although there was transfer of maternal antibodies via egg yolk. The
level of antibodies in egg yolk or hatchling plasma was approximately 10-20% of that in
maternal plasma. Movement of tortoises during relocation, repatriation, or restocking
efforts potentially could transport M. agassizii to previously uninfected sites. Because of
the uncertainty involved in determining latent infections, ELISA-positive animals should
not be moved to locations with no seropositive individuals. However, they can be used in
captive breeding efforts and their offspring released into the wild. Clinically ill animals
should not be relocated to new sites, and should not be housed with clinically healthy
animals in temporary holding situations, such as on-site relocations, or in captive breeding
programs.
IX

CHAPTER 1
INTRODUCTION
Gopher Tortoise Natural History
Gopher tortoises, Gopherus polyphemus, are found in the southeastern United
States, on the coastal plain from southern South Carolina south through Georgia and
throughout Florida, and west through southern Alabama, Mississippi, and Louisiana. The
major population concentrations are in Florida and southern Alabama and Georgia, with
only remnant populations in South Carolina, Mississippi and Louisiana (Auffenberg and
Franz 1982, Diemer 1992a). Populations are concentrated in areas with deep sandy soils
suitable for digging. Vegetation associations in which tortoises are found include longleaf
pine-xerophytic oak woodlands, palmetto scrub, sand pine scrub, oak scrub, beach scrub,
coastal strands, pine flatwoods, dry prairies, native pasture, and savanna, as well as ruderal
habitats (Landers and Speake 1980, Lohoefener and Lohmeier 1981, McRae et al. 1981,
Campbell and Christman 1982, Diemer 1986, Breininger et al. 1988).
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 (Eisenberg 1983).
Gopher tortoises live in loose colonies, with considerable movement of tortoises among
groups over the years (Diemer 1992b). Colonies may be defined more by the availability
of suitable soils for digging burrows, or the distribution of food resources, than by social
1

2
interactions (Campbell and Christman 1982). Gophers are the most fossorial of the four
North American species of tortoises, digging burrows that may extend 5 meters down
from the surface and 15 meters in length (Hansen 1963, Diemer 1986). The burrows
provide a microclimatically stable environment for not only the tortoises, but also for
numerous commensals. Approximately 60 vertebrate speciesfrom snakes to birdsand
over 300 invertebratesincluding spiders, crickets, and beetleshave been found in
tortoise burrows or observed using them as permanent homes or refuges from heat, cold,
fire, and predators (Jackson and Milstrey 1989, Lips 1991, Witz et al. 1991). Some
invertebrate species are obligate commensals, and occur only in active tortoise burrows,
where they feed on tortoise feces and other invertebrates and, in turn, are eaten by other
commensals, such as gopher frogs (Rana areolata) and Florida mice (Podomys
floridanus) (Woodruff 1982). Several species that exclusively or frequently use tortoise
burrows have legal protection in Florida and other parts of their ranges. These include
scarab beetles (F. Scarabaeidae), indigo (Drymarchon coris couperi) and pine (Pituophis
melanoleucus) snakes, gopher frogs, mole skinks (Eumeces egregius), burrowing owls
(Athene cunicularia floridana), and Florida mice (Cox et al. 1987).
The soil disturbance resulting from the digging of the burrows allows deeper
access of air and water into the soil profile, as well as providing bare mineral soil patches
on the surface. When a burrow is abandoned the soil mound, or apron, in front of the
entrance no longer undergoes continual disturbance, allowing certain plants to colonize
the area. The composition of the plant assemblage on the mound may differ from that in
the surrounding undisturbed area, providing a mosaic of small patches in the habitat
(Breininger et al. 1988, McLaughlin 1990). Tortoises consume a wide variety of grasses

3
and forbes, and readily ingest the fruits of many shrubs when available (Gamer and
Landers 1981, Macdonald and Mushinsky 1988). Subsequent tortoise movements may
help spread the seeds to suitable habitat, and some seeds (e.g., gopher apple) may
germinate more readily after passage through the tortoise gut.
Gopher tortoises are long-lived, slow to mature, and have a low reproductive rate.
Age estimates extend up to 100 yr, although 60-80 yr is considered a more reasonable
estimate. Age to sexual maturity ranges from 10 to 20 yr, with possible latitudinal
association (Alford 1980, Iverson 1980, Landers et al. 1982, Wright 1982, Doonan 1986,
McLaughlin 1990, Mushinsky et al. 1994). Gopher tortoises lay one clutch of eggs
annually, generally ranging from 2-10 eggs in size, with the average being 4-8 (Dietlein
and Franz 1979, Iverson 1980, Landers et al. 1980, Linley and Mushinsky 1994). In some
areas with excellent food resources and large tortoises, clutch sizes may average 9-10,
with a range of 6-14 eggs (McLaughlin, 1990). Although some wild individuals have
produced over 20 eggs in one clutch (Godley 1989, L. Macdonald, personal
communication), this is very unusual. With predation rates of up to 95% on eggs and
hatchlings, and further high losses of juveniles aged 1-5 yr, less than 10%, and possibly as
low as 1%, of eggs laid eventually produce reproductive adults (Douglass and Winegamer
1977, Auffenberg and Iverson 1979, Alford 1980, Iverson 1980, Landers et al. 1982,
Diemer 1986, Wilson 1991).
The gopher tortoise is listed in Appendix II of the Convention on International
Trade in Endangered Species of Wild Fauna and Flora (C.I.T.E.S.), which requires
permits for the exportation of the species from the U. S. to any signatory nation, or for re
exportation (Levell 1995). However, if federal or state regulations are more restrictive

4
than C.I.T.E.S., those take precedence. Legal protection is extended to the species in all
states within the range, although the levels of protection vary. The populations west of
the Tombigbee and Mobile Rivers in Alabama, Mississippi and Louisiana are on the
federal threatened species list, which prohibits the taking, exportation, or interstate
movement of individuals originating from that region without permit, and the possession,
transportation, purchase, and/or sale of illegally obtained specimens. Permits issued by the
U. S. Department of Interior can be obtained for scientific research on wild populations,
and possession of limited numbers of individuals for exhibition, education, and/or research.
State permits are required also. The gopher tortoise is fisted in Alabama as a protected
nongame species, in Mississippi as endangered, and in Louisiana as threatened. Louisiana
regulations also prohibit the use of gasoline, chemicals, or volatile substances to flush
reptiles from burrows or other hiding places. Georgia fists the species as threatened and
issues scientific collection permits only to qualified institutions and individuals for
educational or research purposes. Although the law states that burrows may not be
disturbed nor destroyed, nor may explosives, chemicals or smoke be introduced into them
to drive out wildlife, the code explicitly exempts poisonous snakes. Because venomous
snakes, particularly rattlesnakes, use gopher tortoise burrows, tortoises can be adversely
impacted by such activities. South Carolina was the last state to extend legal protection to
gopher tortoises (Mann 1990, Levell 1995), and now fists the species as endangered, with
permits required for any activities involving tortoises. In Florida, where the gopher
tortoise is fisted as a species of special concern (Wood 1996), regulations prohibit taking
and disturbing of tortoises and their habitats, although exceptions are granted regularly for
development, agriculture, and mining operations. In the past, human consumption of

5
tortoises was a major impact on regional distribution, and current poaching activities may
extirpate local aggregations (Diemer 1989, Mann 1990). Large scale conversion of long-
leaf pine habitats to slash pine plantations, transformation of native pasture or savanna to
improved pasture, citrus groves, or row crops, mining operations such as phosphate,
mineral sand, and gravel mines, and urban/suburban development are the main threats to
continued gopher tortoise survival today (Diemer 1986, Cox et al. 1987, Diemer and
Moore 1994). Lack of natural fire regimes due to suppression efforts by humans may alter
vegetation mosaics in remaining habitat, rendering them less suitable to continued
maintenance of tortoise populations (Mushinsky 1986, Mushinsky and Gibson 1991).
Although tortoise numbers increase with increasing areal extent of available
habitat, densities remain constant or decrease (Mushinsky and McCoy 1994).
Fragmentation of mainland areas, with ever smaller islands of suitable habitat surrounded
by agricultural and urban development, may force tortoises into higher density populations
than would occur normally. Vegetational changes resulting from reduced fire incidence in
small areas, particularly increased canopy closure and decreased herbaceous vegetation
(Mushinsky 1985, 1986), can lead to decreases in reproductive rates and juvenile survival
(Aufifenberg and Franz 1982, McLaughlin 1990, Mushinsky and McCoy 1994). Increased
intraspecific interactions at higher densities (McRae et al. 1981) may lead to elevated
physiological stress, potentially affecting immune system function and rendering animals
more susceptible to disease. Increased densities and number of interactions multiply the
opportunities for transmission of communicable agents, making fragmented populations
more likely to sustain high morbidity during epizootics. If mortality is high, lowered
reproduction due to habitat degradation may be insufficient for population recovery.

Along with increased physiological stress placed on tortoise populations from
human activities, there may be toxicological or immunological stress from chemicals
introduced into the environment. No research has been conducted into the effects of
herbicides, fungicides, insecticides, and fertilizers on gopher tortoise health, growth, or
reproduction. Investigations of disease in free-ranging populations of tortoises have
begun only recently.
Tortoises and Upper Respiratory Tract Disease
Although individual captive and wild gopher tortoises have been observed with
clinical signs of respiratory diseases for over 20 yr (E. R. Jacobson, unpublished data), the
first documentation of a larger-scale disease outbreak was in 1989, when an epizootic of
upper respiratory tract disease (URTD) was documented on Sanibel Island, Lee County,
Florida (G. S. McLaughlin and M. Elie, unpub. data). With the loss of 25-50% of
breeding-age adults in one population, recovery could take 50-150 yr (G. S. McLaughlin
unpub. data), barring further major losses and without substantial habitat improvement
leading to increased recruitment.
In the 1980s, large-scale population reductions (33-76% over 10 yr) of desert
tortoises (Gopherus agassizii) were documented at several sites in the western Mojave
Desert of California and at one site in the eastern Mojave (Com 1994, Berry in press).
Tortoises with clinical signs of URTD were observed among the remaining populations at
several sites (Knowles 1989; Berry 1990, in press). As a result of the declines, tortoises in
the Mojave Desert north and west of the Colorado River were declared threatened (U.S.
Fish and Wildlife Service 1990).

7
Clinical signs of URTD in gopher and desert tortoises include serous, mucoid, or
purulent discharge from the nares, excessive tearing to purulent ocular discharge,
conjunctivitis, and edema of the eyelids and ocular glands (Jacobson et al. 1991, G. S.
McLaughlin personal observations). Individual infected tortoises vary in the suite of signs
they have, and the severity can vary from day to day. Nares may become occluded with
caseous exudate, preventing externally visible nasal discharge. Lymphocytic infiltration of
the comeas, while rare, may decrease an animals ability to forage or avoid predators.
Tortoises may become lethargic and anorectic, leading to dehydration, emaciation, and
eventual death from cachexia. Lethargy, nonresponsiveness to stimuli, and altered
behavior pattemssuch as basking at lower temperatures than normalmay render a
tortoise more susceptible to predation. Moribund animals often develop petechial to
ecchymotic hemorrhages under the scutes, especially visible on the plastron (G. S.
McLaughlin pers. obs.), which may be due to septicemia caused by secondary infection
with opportunistic bacteria.
Several agents were hypothesized to cause respiratory tract disease in tortoises,
including viruses, Mycoplasma sp. (Lawrence and Needham 1985), and Pasteurella
testudinis (Snipes and Biberstein 1982). However, experimental infections to determine
the etiologic agent were not conducted. Beginning in 1989, efforts were undertaken to
determine the etiology of URTD. Clinically ill and healthy desert tortoises from California
were examined and samples collected, either from live animals or at necropsy.
Hematologic and serum biochemical evaluations, liver vitamin and metal determinations,
and pathologic and microbial investigations were conducted (Jacobson et al. 1991).

8
Hemoglobin concentration and phosphorous levels were lower in ill than in healthy
tortoises, and ill tortoises had higher sodium, blood urea nitrogen (BUN), creatinine,
serum glutamic oxalacetic transaminase (SGOT) activity, and total cholesterol levels.
Levels of liver and serum vitamins A and E did not differ between healthy and ill tortoises,
nor did liver values of selenium, copper, lead, or cadmium. However, iron and mercury
were higher in ill than in healthy tortoises. Healthy tortoises did not exhibit any
histological abnormalities in nasal mucosa or other tissues. Ill tortoises had less fat than
controls, atrophied thymuses, increased numbers of lymphocytes in the sinusoids of the
spleens, and increased amounts of iron granules in the hepatocytes. The most striking and
consistent changes were in the nasal passage and cavity tissues (Jacobson et al. 1991).
Grossly, passages and cavities of ill tortoises contained moderate to large amounts of
serous to purulent exudate. Histologically, the tissues exhibited loss of mucosal glands
and goblet cells, proliferation of epithelial cells, infiltrates of lymphocytes and histiocytes,
loss of cilia, dysplasia of the olfactory epithelia with infiltration of heterophils, basal cell
hyperplasia, occasional squamous metaplasia, and occasional erosion and ulceration of
mucosal epithelia. By electron microscopy, pleomorphic organisms resembling
Mycoplasma sp. were seen on cell surfaces and tightly adhered to cell membranes of ill
tortoises, but not healthy ones. Significantly more Pasteurella testudinis were isolated
from ill than healthy tortoises, and Bacillus sp. and Mycoplasma-like organisms were
isolated only from ill tortoises. By electron microscopy, the latter resembled those seen on
the nasal cavity tissues of ill tortoises (Jacobson et al. 1991).
The mycoplasma was determined to be a new species, provisionally named
Mycoplasma agassizii (Brown et al. 1994). An enzyme-linked immunosorbent assay

9
(ELISA) to detect antibodies against the mycoplasma in plasma and serum samples was
developed (Schumacher et al. 1993), and experiments were undertaken to fulfill Kochs
postulates. The disease was induced by inoculation of tortoises with pure cultures of the
mycoplasma, but not Pasteurella testudinis (Brown et al. 1994). Histologically, the
lesions were consistent with those seen in the previously examined naturally infected
tortoises. Additional work led to the development of a polymerase chain reaction (PCR)
test to detect the bacteria in nasal flush and swab samples (Brown et al. 1995).
Histologic examination of nasal tissues and microbiologic evaluation of flush and
swab samples from clinically healthy and clinically ill tortoises from Las Vegas Valley,
Nevada, resulted in findings in the ill tortoises similar to those in the California tortoises,
with 92% showing lesions of URTD, 50% being culture positive for M. agassizii, and
100% reacting positively in the ELISA (Jacobson et al. 1995). However, 73% of the
healthy tortoises had lesions consistent with URTD, 50% were culture positive for M
agassizii, and 42% were seropositive for antibodies against the bacteria. These findings
demonstrate that the disease can exist in a subclinical form in a substantial proportion of a
population. An annual cycle of convalescence and recrudescence of clinical signs has been
seen in captive desert (I. M. Schumacher, pers. comm.) and gopher (D. L. Morris, pers.
comm.) tortoises. Other mycoplasmal diseases also can exist as chronic, subclinical
infections, with recurrence of clinical signs and increases in transmission potential when
the host is stressed (see below). Annual fluctuations in temperature, rainfall, and forage
availability may be sufficient to cause detectable outbreaks in an infected population.
Increased morbidity and mortality may occur in times of unusually severe environmental
stress, such as prolonged drought, hurricanes, excessive rainfall with flooding of burrows,

10
or very cold winters. Human impacts on tortoises and their habitat, whether through
disruption of normal behavior patterns, degradation of habitat through agriculture,
silviculture, mining or development operations, or pollution, may cause sufficient
physiological stress to trigger proliferation of the mycoplasma and recurrence of signs.
Capturing and transporting of tortoises during relocation, restocking and repatriation
efforts also may be significant sources of stress.
Mycoplasmal Respiratory Diseases in Domestic Animals and Humans
Mycoplasmas cause respiratory disease in other taxa, including rodents, poultry,
swine, ruminants, and humans. All have similar pathological effects, which are described
below, and many are exacerbated by concurrent infections or environmental stress.
Murine respiratory disease caused by Mycoplasma pulmonis has caused problems
in laboratory settings for more than 70 yr (Lindsey et al. 1971). It has necessitated
complex and expensive control measures, including various types of isolation mechanisms
and cesarean delivery of mice and rat pups to reduce the prevalence of M. pulmonis
infections in colonies (Cassell et al. 1984, Davis et al. 1987). Interactions among
environment, host, and strain factors influence the impacts at individual and population
levels (Simecka et al. 1992). Histologically, lesions are characterized by lymphoid
hyperplasia and chronic inflammation (Cassell et al. 1985).
Poultry respiratory diseases can be caused by viruses, mycoplasmas, and other
bacteria, singly or in combination. Without concurrent viral or other bacterial infections,
disease can be mild or subclinical (Kerr and Olson 1967). Environmental factors,
particularly dust and ammonia levels, as well as strain differences, affect the severity of

11
outbreaks of mycoplasmosis. Other stress factors, such as crowding and cold weather,
also influence morbidity and mortality (Jordan 1972). Mycoplasma gallisepticum causes
chronic respiratory disease in chickens and sinusitis in turkeys, and affects ring-necked
pheasants (Phasianus colchicus), chukar (Alectoris chukar), northern bobwhite (Colinus
virginianus), common peafowl (Pavo cristatus) (Yoder 1991) and Japanese quail
(Coturnix japnica) (do Nascimento and do Nascimento 1986). Lower respiratory tract
complications are seen rarely in turkeys (Simecka et al. 1992). Histologically, hypertrophy
and hyperplasia of respiratory epithelia, including mucous cells, are seen, as is lymphoid
infiltration of the submucosa. Heterophilic exudate is seen in the airways (Nunoya et al.
1987, Trampel and Fletcher 1981). Mycoplasma meleagridis infection is seen primarily in
chicks and poults up to 10 wks of age, and is sexually transmitted. Turkeys, Japanese
quail, and peafowl develop air sacculitis, sometimes with accompanying tracheitis and
pneumonia, but not sinusitis. Histologically, the lesions are characterized by hypertrophy
and hyperplasia of the air sac epithelia, with edema and lymphoid infiltration (Stipkovits
1979). Young chickens and turkeys also are susceptible to M. synoviae, usually in
conjunction with Newcastle disease virus or infectious bronchitis (Hopkins and Yoder
1984, Springer et al. 1974). As with M. gallisepticum, turkeys develop sinusitis
(Stipkovits 1979). All three bacteria can be transmitted vertically (i.e., via the egg)
(Simecka et al. 1992).
Swine develop mild pneumonia when infected by M. hyopneumoniae, and although
mortality is virtually nil, the disease is chronic, causing slow growth and reduced weight
gains, decreasing profitability (Oboegbulem 1981, Jericho 1986). Although signs can
disappear, recurrences will occur with weather changes, viral infections, and other

12
stressors (Whittlestone 1976). Lesions are alveolar, initially characterized by neutrophilic
infiltrates and later by lymphocytes and macrophages (Baskerville 1972).
Contagious bovine pleuropneumonia (CBPP), although not present in the United
States, is an economically important disease of cattle and water buffalo caused by M
mycoides subsp. mycoides (Howard and Gourlay 1978, Trichard et al. 1989). Morbidity
rates can reach 100% with up to 50% mortality, and recovered animals maintain a carrier
state. Neutrophilic exudate in the airways, serous exudate in the pleural cavity,
serofibrinous exudate in alveoli, and edema and necrosis of regional lymph nodes and
interlobular septa are seen at necropsy. Interseptal, peribronchial, and perivascular
lymphoid infiltration can be seen histologically (Hudson 1971). Calf pneumonia, with high
mortality among dairy calves, is caused primarily by viruses, but can be caused by either
M bovis or M. dispar (Stalheim 1983, Bryson 1985). Infection with the former is
characterized by peribronchiolar and alveolar monocyte infiltration, while the latter causes
interstitial pneumonia with monocyte infiltration of alveolar walls, but not peribronchiolar
infiltration. Mycoplasma dispar also causes superficial and asymptomatic infection of
respiratory mucosa (Woldehiwet et al. 1990).
Lambs are susceptible to infection with M. ovipneumoniae, which is transmitted to
neonates from the ewes. Pneumonia, characterized by coughing, sneezing, nasal
discharge, fatigue, and poor weight gain, can develop as colostral antibodies wane.
Lesions are characterized by alveolar proliferation, nodular lymphoid hyperplasia, and
peribronchial and perivascular lymphoid infiltration (Carmichael et al. 1972, Foggie et al.
1976). Severity can be exasperated by infection with Pasteur ella haemolytica biotype A
(Jones et al. 1982).

13
Goats can suffer 60-100% mortality when infected by M. mycoides subsp.
mycoides, M. mycoides capri, or Mycoplasma strain F38, making contagious caprine
pleuropneumonia the most economically important goat disease. Pathologically, signs and
lesions are similar to those seen in CBPP, with the addition of polyarthritis (Cottew 1984).
Humans are susceptible to infection with M. pneumoniae, which causes
tracheobronchitis and, less commonly, primary atypical, or walking, pneumonia (Chanock
et al. 1963, Clyde 1983). The upper respiratory tract is affected, not the alveoli, and otitis
media is seen also (Clyde 1979, Mansel et al. 1989). Immunopathological sequelae,
although rare, can include arthritis, dermal lesions, cardiopathy, and neurological
complications (Johnston et al. 1983, Lyell et al. 1967, Naftalin et al. 1974, Murray et al.
1975). Histologically, perivascular and peribronchial monocytic infiltrations with some
neutrophilic exudate are seen (Dajani et al. 1965). Most commonly affected are 5-9 yr old
children, with a decline in incidence until about age 25, then an increase in 30-40 yr olds
(Denny et al. 1971). The disease is endemic, with slight increases in late summer and fall,
and cyclic epidemics occurring at 4-7 yr intervals (Krause and Taylor-Robinson 1992).
Mycoplasmal Diseases of Wildlife
Research on the impact of mycoplasmosis on wildlife is limited, but recent
developments are provoking interest. In early 1994, house finches (Carpodacus
mexicanus) in the mid-Atlantic and northeastern regions of the United States were found
with conjunctivitis, rhinitis, and sinusitis. Many birds were found dead or dying and were
submitted to wildlife care facilities, veterinary hospitals, and state agricultural laboratories.
Many of those birds had lesions that were histologically compatible with mycoplasma

14
infections, and were infected with M. gallisepticum (Ley et al. 1996, Luttrell et al. 1996).
Since then, the organism has been associated with morbidity and mortality in other species
of passerines, including goldfinches (Carduelis tristis) (Nettles 1996), and was transmitted
to a blue jay (Cyannocitta cristata) in a rehabilitation facility (Ley et al. 1996).
Conjunctivitis and M gallisepticum also were found in house finches in Georgia (Luttrell
et al. 1996). The potential for this organism to spread over a large area due to the long
distances traveled by migratory birds, the mixed species flocks in which they congregate,
and local concentrations of many species around bird feeders is of great concern to many
ornithologists and ecologists, as well as poultry producers. There are indications that the
strain, while highly pathogenic to chickens, is not readily transmissible to poultry under
natural conditions (Nettles 1996). A new species of mycoplasma (M sturni) was isolated
from the conjunctiva of a European starling (Sturua vulgaris) with conjunctivitis found in
Connecticut during the epomitic (Forsyth et al. 1996). Although it was isolated in pure
culture, it was described as growing rapidly, whereas pathogenic mycoplasmas typically
grow slowly; therefore, it could have overgrown M. gallisepticum if that species had been
present. The pathogenicity of M. sturni needs to be investigated.
Three new species of mycoplasmas have recently been described from raptors in
Europe (Poveda et al. 1994). All were associated with respiratory diseases clinically and
histologically consistent with those caused by Mycoplasma spp., with lesions including
hyperplasia of mucous glands, lymphoid hyperplasia, and perivascular cuffing (Poveda et
al. 1990). The species were isolated from buzzards (Buteo buteo), saker falcons (Falco
cherrug), and griffon vultures (Gyps fulvus), and have been named, respectively, M.
buteonis, M. falconis, and M. gypis. Pathogenicity of these species, their distribution in

15
wild, free-ranging birds, and their potential impacts on the populations need to be further
investigated.
Several species of mycoplasmas, including M. cloacale (Bradbury et al. 1987,
Goldberg et al. 1995) and M. anatis (Ivanics et al. 1988, Poveda et al. 1990, Goldberg et
al. 1995), have been isolated from semi-domestic and wild ducks and other avian species
throughout the world. Stipkovits et al. (1986) reported isolation ofM cloacale from
geese with inflammation of the cloaca and phallus, but Goldberg et al. (1995) found no
association ofM cloacale with disease in wild mallards (Anas platyrhynchos), black
ducks (A. rubripes), or canvasbacks (A. valisneria). Stipkovits (1979) reported
pathogenicity of M. anatis to domestic ducklings and eggs, and neurological signs have
been recorded in ducks infected with M. anatis (Ivanics et al. 1988). Samuel et al. (1995)
infected game-farm mallard eggs with M. anatis and found reduced hatchling success,
hatchling size and growth rates. Hatchlings infected at 1 d of age did not have slower
growth rates. Goldberg et al. (1995) found esophagitis, tracheitis, and vaginitis in female
mallards from which M. anatis was isolated, and presented evidence for vertical
transmission ofM anatis in a wild gadwall (Anas strepera). Potentially, M anatis
infections could reduce recruitment in wild duck populations. Experimental infection of
ducklings withM gallisepticum resulted in suppressed growth rates (Stipkovits 1979).
Four other unidentified mycoplasmas were isolated from mallards, gadwalls, and black
ducks, but could not be associated definitively with disease (Goldberg et al. 1995).
Although M gallisepticum has been isolated from wild turkeys, most cases have
occurred in birds with close association to domestic poultry (Davidson et al. 1982, Jessup
et al. 1983, Luttrell et al. 1991, Fritz et al. 1992). Even though experimental infection of

16
pen-reared wild turkeys has resulted in decreased productivity (Rocke et al. 1988), there is
little evidence to indicate that infection with M. gallisepticum strains commonly occurring
in domestic fowl poses a threat to wild turkey populations (Luttrell et al. 1991), or that
wild turkey populations are important in the epizootiology ofM gallisepticum (Davidson
et al. 1988). Mycoplasma gallopavonis has been isolated from wild turkeys in Texas
(Rocke and Yuill 1987), South Carolina, Georgia (Luttrell et al. 1991), Colorado, New
Mexico, and Oklahoma (Fritz et al. 1992). Although lethal to experimentally infected
domestic turkey eggs (Rocke and Yuill 1987), the pathogenicity of M. gallopavonis to
wild turkeys has not been investigated. Mycoplasma synoviae and other, untyped,
Mycoplasma spp. were isolated from turkeys in Arizona (Fritz et al. 1992), but no
association with disease was found. As with M. gallisepticum, wild turkeys do not appear
to be important in the epizootiology of M. synoviae or M. me/eagridis (Davidson et al.
1988).
In 1993 an epizootic of polyarthritis occurred in juvenile farmed crocodiles
(Crocodylus niloticus) in Zimbabwe. The outbreak was characterized by high morbidity,
but low mortality. A mycoplasma was isolated, determined to be a previously
unrecognized species, and named M. crocodyli (Mohan et. al 1995). In 1995 a die-off of
captive adult American alligators (.Alligator mississippiensis) at a private facility in
Florida, with veiy high mortality, was associated with systemic infection with a different
species of mycoplasma, also previously unrecognized (Brown et al. 1996a). The name M.
lacerti has been proposed. In addition to M. agassizii, a second mycoplasma was found in
desert tortoises with evidence of URTD, including clinical signs, histologic lesions, and/or
positive ELISA tests (Brown et al. 1995). In a small pilot study, that organism was found

17
to cause disease and seroconversion in gopher tortoises previously unexposed to either
mycoplasma (D. R. Brown, et al., unpub data).
Mycoplasmas have been isolated from bighorn sheep (Ovis canadensis) with
pneumonia associated with P. haemolytica, and there is concern that these may have a role
in the disease process (D. Hunter, pers. comm.). A mycoplasma isolated from dead and
live captive bighorn sheep with pneumonia was typed asM arginini (Woolf et al. 1970).
Apparently, no further work has been done regarding the pathogenicity of that strain to
bighorn. With continued exposure of bighorn to domestic livestock, Mycoplasma spp.
could interfere with recovery efforts. An outbreak of pneumonia associated with M.
ovipneumoniae infection occurred in captive Dalis sheep (Ovis dalli dalli) following
indirect exposure to domestic sheep (Black et al. 1988). Although Dalis sheep are
unlikely to be exposed to domestic livestock in their native habitat, it is clear that captive
herds or those in transit must be protected from exposure to pathogens.
Behymer et al. (1989) detected antibodies to Mycoplasma spp. in mule deer
(Odocoileus hemiotms) in California, but there was no association with disease, and
isolations were not attempted. One-humped camels (Camelus dromedarius) and African
buffalo (Syncerm caffer) on a game farm in Kenya had antibodies against Mycoplasma
strain F38, and camels were seropositive for M. mycoides mycoides, but no isolations
were made and no disease was seen (Paling et al. 1988). Kirchoff et al. (1996) found, in a
survey of captive arthritic elephants (Elephas maximus and Loxodonta africana), that
about 60% of the females harbored a new mycoplasma in the urogenital tract, although
none was recovered from males. It is not known if the mycoplasma, named M. elephantis,
causes arthritis.

18
Chronic Manifestations of Mycoplasmal Infections
Many mycoplasmal diseases are characterized by an overaggressive or
inappropriate immune response by the host, eventually leading to autoimmune damage to
the affected sites, whether respiratory or urogenital tract, joints, heart, skin, or other organ
systems (Krause and Taylor-Robinson 1992, Simecka et al. 1992, Cole 1996). Infected
hosts may be more susceptible to secondary infections with other bacteria or viruses. In
wildlife species, such complications may reduce the fitness of the animals by altering
behavior, leading to decreased foraging efficiency, increased susceptibility to predators, or
diminished mate seeking behavior. Energy that normally would be allocated to
reproduction may be needed to repair or compensate for damage to multiple organ
systems. Therefore, even if mycoplasmal infections do not cause mortality directly, they
can affect individual and population viability.
Project Overview and Specific Objectives
Due to the listing by the Florida Game and Fresh Water Fish Commission of the
gopher tortoise as a species of special concern, and the subsequent permitting of over 450
relocations involving more than 8000 tortoises, particular attention has been focused on
the dynamics and persistence of both natural and relocated populations. Understanding
the effects of URTD on individuals and populations is essential for proper management of
remaining populations; therefore, a study was begun in 1993 on the etiology, pathology,
and diagnosis of URTD in gopher tortoises. The original objectives were 1) to describe
the pathology of natural infections and identify possible etiologic agents, 2) to perform

19
experimental infections to determine conclusively the etiologic agent and dose-related
effects, 3) to refine the PCR test and delineate further the taxonomy of the tortoise
mycoplasmas, 4) to characterize the immune responses of gopher tortoises to URTD
pathogens, and 5) to compare the efficacy of culture, PCR and serology in detecting new
infections.
As the original experiments were conducted and concern over the continued well
being of tortoise populations throughout the range increased, additional questions were
raised. Those were concerned primarily with management and conservation of gopher
tortoises, particularly with respect to relocation, repatriation, and restocking efforts.
Several questions were raised repeatedly in meetings with state and federal agency
personnel, particularly regarding possible immunity to the organism and factors related to
transmission. In order to address those questions additional objectives were established,
as delineated in specific objectives 2-7 below, and discussed in Chapters 3 through 8.
Chapter 2 will detail the methods common to fulfilling the objectives.
This dissertation addresses the following specific objectives:
1) To describe the clinical presentation and lesions of natural infections by M. agassizii in
gopher tortoises and compare the lesions to those found in desert tortoises (Chapter 3).
2) To test the hypothesis that tortoises that have produced antibodies against M. agassizii
are protected against reinfection with the organism and subsequent development of
URTD (Chapter 4).
3) To assess the probability of horizontal transmission of M. agassizii between adult
tortoises and to test the hypothesis that transmission is more likely to occur from an

20
infected host that is clinically ill and PCR or culture positive than from a seropositive
host that is not clinically ill or is clinically ill but culture or PCR negative (Chapter 5).
4) To determine ifM agassizii is transmitted vertically (Chapter 6).
5) To assess the relationship of antibodies in eggs and hatchling serum to those in
maternal serum (Chapter 6).
6) To collect preliminary data to test the hypothesis that M. agassizii can be transmitted
environmentally (e.g., in burrows) (Chapter 7).
7) To discuss the implications of the findings of the above research and additional
concurrent research on conservation and management of gopher and other tortoises
(Chapter 8).

CHAPTER 2
METHODS
Tortoises. Intake Procedures. Clinical Assessments and Sampling Methods
For the natural infection studies, tortoises were obtained from various locations in
the state of Florida under Florida Game and Fresh Water Fish Commission permits
number WX93227 issued to Elliott R. Jacobson and number WX94037 issued to Mary B.
Brown. Tortoises were processed the day of arrival at the University of Florida (UF),
Gainesville. For the natural and experimental infection studies, gopher tortoises were
transferred from a development site in central Florida to UF in April, July, and August
1994 and April 1995, and processed the day following arrival. Tortoises were examined
for clinical signs of URTD: nasal and ocular discharge, palpebral edema, and
conjunctivitis. The signs were graded individually on a scale of 0-3, none to severe. Total
clinical sign score was calculated as the nasal discharge score plus the mean of three
ocular sign scores (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. Straight line
carapace length, thickness, and width were measured to the nearest cm with forestry
calipers.
21

22
A 2-3 ml blood sample was drawn from the jugular or brachial vein and placed in a
lithium heparin Vacutainer tube (Becton Dickinson and Company, Rutherford, New
Jersey). Blood was centrifuged and an aliquot of plasma was removed for antibody
screening by an enzyme-linked immunosorbent assay (ELISA) (see below).
After cleansing the area around the nares with alcohol dampened gauze, nasal flush
samples were collected by flushing with approximately 0.5 ml sterile SP4 broth using a 1
ml syringe without needle. Calcium alginate-tipped swabs were inserted gently into the
nares, and a sample was obtained and streaked onto SP4 agar plates.
Culture Procedures
A 100 pi aliquot of the flush sample was used for polymerase chain reaction (PCR)
analysis; the remaining sample was serially diluted ten-fold to 102 and incubated at 30C
for a maximum of three weeks, or until determined to be positive or contaminated.
Twenty pi of each dilution were placed on SP4 agar and incubated at 30C and 5% CO, as
were the streak plates. Plates were examined regularly for a maximum of six weeks to
detect the growth of mycoplasma. In the second year of the study, broth cultures were
incubated for 24-48 hr before removing the aliquot for PCR, as that modification increased
the sensitivity of the PCR (G. S. McLaughlin and D. R. Brown, unpub. data).
PCR Procedure
Nasal aspirate samples were analyzed for the presence of Mycoplasma agassizii
DNA based upon PCR amplification of the 16S rRNA gene (Brown et al. 1995). Nasal
flush and culture samples were centrifuged for 60 min at 14,000 rpm at 4C, and the

23
supernatant aspirated. Three to four microliters of 20 pg/ml proteinase K (Sigma, St.
Louis, MO) in 20 pi lysis buffer (100 mM tris pH 7.5, 6.5 mM DTT, 0.05% Tween 20)
were added to the pellets, which were resuspended, and the samples were incubated at
37C for 8-16 hours. After denaturing the proteinase K at 97C for 15 min, 5 pi of each
sample were removed and added to 45 pi of reaction solution containing two primers for
the 16S rRNA gene at 1 pM each, deoxynucleoside triphosphates at 200 pM, 2.0 mM
MgCl, and 2.5 units of Taq polymerase (Promega, Madison, WI). 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', Brown et al.
1995) and a Mycoplasma genus-specific region [anti-sense strand nt 1055 to 1031,
5'-TGCACCATCTGTCACTCTGTTAACCTC-3', Van Kuppeveld 1992], Samples were
subjected to 50 cycles of template denaturation for 45 sec at 94C, primer annealing for 1
min at 55C, and polymerization for 45 sec at 72C, followed by 10 min at 72C. Positive
samples yielded 576 base pair (bp) products that were visualized by combining 15 pi of
product with 2 pi bromphenol blue in 50% glycerol solution and electrophoresing on
ethidium bromide-stained 1.5% agarose gels in tris-borate-EDTA buffer. Positive control
samples using 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, Hind III digest of x phage DNA, was included on each gel.
In order to confirm that the isolates obtained from naturally and experimentally
infected tortoises were M. agassizii, an additional procedure, restriction fragment length
polymorphism (RFLP) analysis, was conducted on at least one isolate from each tortoise.
Twenty microliter samples of products from the above amplification procedure were

24
incubated with 10-20 units of the endonuclease Agel (New England Biolabs, Inc.,
Beverly, MA), which cuts the M agassizii amplification product at nt 613, and 5 pi of
reaction buffer, at 25C for one hour, and the products electrophoresed as above. The
procedure resulted in products of 434 and 142 bp from M. agassizii-positive samples, and
no change in non-M agassizii- samp les.
ELISA Procedure
An aliquot of plasma from each sample was used for determinations of levels of
antibodies specific for M agassizii (Schumacher et al. 1993). Ninety-six-well microtiter
plates (Maxisorp F96, Nunc, Kamstrup, Denmark) were coated with 50 pi of a whole-cell
lysate of M. agassizii strain 723 at 10pg/ml in phosphate buffered saline with 0.02% azide
(PBS-AZ). Plates were incubated overnight at 4C, 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 4C with 250 pl/well PBST containing 5%
non-fat dry milk (PBS-TM). Following washing, 50 pi of plasma diluted appropriately for
the specific study with PBS-TM were added to individual wells in duplicate or triplicate,
and the plates were incubated at room temperature for 60 min. The plates were washed,
50 pl/well of a biotinylated monoclonal antibody (MAb HL673) against the light chain of
desert tortoise immunoglobulins IgY and IgM at 1 pg/ml in PBS-TM was added, and
plates were incubated for 60 min. Following washing, a conjugate of alkaline phosphatase
and streptavidin (AP-S; Zymed Laboratories, Inc., San Francisco, CA) at 1:2000 in PBS-
AZ was added at 50 pl/well, plates were incubated for 60 min, and washed. Substrate, p-
nitrophenyl phosphate disodium (pNPP; Sigma), was prepared at 1 mg/ml in 0.01 M

25
sodium bicarbonate, pH 9.6, with 2 mM MgCl2, and added to wells at 100 pi/well. Plates
were incubated for 60 min in the dark, then read at 405 nm on a microplate reader (EAR
400 AT, SLT Labinstruments, Salzburg, Austria). The mean of two or three wells coated
with antigen and incubated with conjugate and substrate only was used as the blank. A
positive control, plasma from a naturally infected gopher tortoise from Sanibel Island, and
a negative control, plasma from an uninfected tortoise from Orange County, were included
on each plate.
Results from the ELISA were optical density (OD) readings from the microplate
reader. The OD readings reflected the intensity of the yellow color developed when all
components of the reaction (specific tortoise antibodies against M. agassizii, biotinylated
MAb HL673, AP-S, and /?NPP) were present. The readings are on a continuous scale, but
were interpreted categorically by calculating the ratio of the sample readings to the
negative control reading. Ratio values less than or equal to 2.0 were considered negative,
those greater than 2.0 and less than or equal to 3.0 were classified as suspect, and those
greater than 3.0 were classified as positive.
Study Group Assignment
Tortoises exhibiting one or more signs of disease, or with a positive culture or
PCR result, were designated as diseased. Animals were designated clinically healthy if
found free of signs of URTD, and with negative culture and PCR results. Tortoises were
assigned to study groups based on sex, weight, clinical assessments, culture, PCR and
ELISA results, and placed in the appropriate pens.

26
Husbandry Procedures
Tortoises were housed singly or in pairs in outdoor pens at the UF Animal
Resource Farm (ARF). There were four groups of 10 pens in a larger, chain link-fenced
enclosure (Fig. 2-1). Pens were approximately 21 m2, constructed of a wooden frame
with sheet metal extending vertically approximately 0.7 m above and below ground,
partially covered by shade cloth, and provided with an artificial burrow, a water dish, and
a cement feeding stone. Pens were observed daily by ARF staff, and watered daily in the
summer and as needed in the winter. The tortoises were fed a salad of mixed vegetables
three times per wk, and fruit was provided on an occasional basis. In addition, I observed
the pens and tortoises from three to seven days each week, with some days including
multiple observations. Because of individual behavior patterns, not every tortoise was
observed at each time point. The amount of food eaten was recorded for each pen the day
following feeding, and the stones were cleaned. Remaining food was collected and bagged
separately for each section, using brushes and dust pans assigned to that section. Stones
were then sprayed with a dilute (1:30) bleach solution, allowed to soak for a few minutes,
and hosed off. Water dishes were rinsed and filled daily, and bleached and scrubbed as
necessary. 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 a different tortoise. Used gloves were placed in plastic garbage bags and
disposed of properly. Before entering the first pen on a given day, personnel sprayed their
footwear with a dilute bleach solution. Footwear was sprayed with bleach before leaving

27
a pen. Animals were captured by hand or using wire cage-type traps (Tomahawk Live
Trap Company, Tomahawk, WI) that were covered with brown paper to protect the
animals from the weather. Traps were cleaned, sprayed with bleach solution, and allowed
to air dry following each use. 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, WI) for transport and holding. Containers were
bleached, scrubbed, and washed in an automatic cage washer before reuse.
D C
A B
Figure 2-1. Layout of tortoise pens at the University of Florida Animal Resource Farm.
The outer fence was chain link, and the inner fence and pen dividers were corrugated sheet
metal on a wooden frame.
Necropsy Procedures
All diseased and selected healthy tortoises were euthanatized with a combination of
drugs. Ketamine was administered intramuscularly at 60 80 mg/kg followed by a

28
concentrated barbiturate solution (Socumb, The Butler Company, Columbus, Ohio, USA)
intraperitoneally at 1 ml/kg. Once the tortoises showed complete muscle relaxation and
were unresponsive to painful stimulation, they were exsanguinated via a 23 gauge butterfly
catheter inserted into the carotid artery and then decapitated. Flush and swab samples
were collected as previously described, then the head was bisected longitudinally with an
electric saw. Following bisection, the cartilage over each nasal cavity was reflected
aseptically, and flushes and swabs of 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 and the following tissues were collected, fixed in neutral buffered 10%
formalin, sectioned at 5-6 pm, and stained with hematoxylin and eosin: glottis, cranial
trachea, tracheal bifurcation, left lung, right lung, thyroid, heart, brain, thymus, esophagus,
stomach, small intestine, pancreas, large intestine, cloaca, spleen, liver, left and right
kidney, bladder, right and left gonads, chin gland, buccal salivary gland, chin gland, and
tongue. Tissues were examined by light microscopy and abnormalities or changes were
recorded.
Histopathology Procedures
For histopathologic studies, heads were fixed in 10% neutral buffered formalin
(NBF), decalcified, embedded in paraffin, sectioned longitudinally at 5-6 pm, and stained
with hematoxylin and eosin. Sections were examined by light microscopy and classified on
a scale of 0 to 5, with 0 being normal and 5 exhibiting severe inflammation and / or
changes. Changes in the epithelium and submucosa were recorded separately.

29
The following criteria were utilized for grading lesions:
Normal (score = 0): Occasional small subepithelial lymphoid aggregates; rare
heterophils in the lamina propria. No changes in mucosal or glandular epithelium; no
edema.
Mild (1): Multifocal small subepithelial lymphoid aggregates; multifocally, small
numbers of heterophils, lymphocytes, and plasma cells in the lamina propria; mild edema in
lamina propria; minimal changes in mucosal epithelium.
Moderate (2-3): 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; proliferation and disorganization of the basal epithelium.
Severe (4-5): Focally extensive to diffuse band of lymphocytes and plasma cells
subjacent to and obscuring the overlying mucosal epithelium; large numbers of heterophils
in lamina propria and infiltrating 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; occasional squamous metaplasia.
Statistical Analyses
Statistical analyses were performed using SAS (SAS Institute, 1988) or SigmaStat
for Windows, Version 1.0 (Fox et al. 1994). Because the analyses varied for each
experiment, the specific methods will be addressed in the appropriate chapters.

CHAPTER 3
NATURALLY OCCURRING UPPER RESPIRATORY TRACT DISEASE
Methods
Tortoises
Twenty-three gopher tortoises from the following locations in Florida (Figure 3-1)
were transported to UF from August 1993 to September 1995: Alachua County (n = 2),
Sanibel Island, Lee County (n = 3), Volusia County (n = 1), St. Lucie County (n= 1),
Indian River County (n = 1), Orange and/or Osceola Counties (n = 15). Collection of
tortoises, except those from Orange and Osceola Counties, was opportunistic, and a result
of submissions to the UF Wildlife Clinic, or other veterinary clinics. Some tortoises had
exhibited signs of URTD, while others had been hit by automobiles. Tortoises from
Orange and Osceola Counties were selected on the basis of clinical evaluations, ELISA,
culture, and / or PCR results. Six clinically healthy animals were included in the latter
group. Tortoises were evaluated for clinical signs of URTD (i.e., nasal and ocular
discharge, palpebral edema, and conjunctivitis) and those exhibiting one or more signs of
disease, with a past history of clinical signs, with positive culture or PCR results, or with a
positive ELISA result, were designated as diseased. Tortoises were designated healthy if
they were free of any history of or current signs of URTD and were culture, PCR, and
ELISA negative.
30

31
Figure 3-1. Locations in Florida from which gopher tortoises were obtained. 1 Alachua
County, 2 Volusia County, 3 Orange/Osceola Counties, 4 Indian River
County, 5 St. Lucie County, 6 Sanibel Island, Lee County.
Necropsy and Histology Procedures
Necropsy procedure and light microscopic evaluation of tissues were performed as
detailed in Chapter 2. Gross necropsies were conducted on 21 tortoises. Multiple tissues
were collected from 15 tortoises, heads and livers from six and only heads from two.
Microbial Isolation
Flush and swab samples for Mycoplasma isolation were collected from the nasal
passages and cavities of each tortoise and processed as described in Chapter 2. Swab
specimens of the dorsal nasal cavities of 16 tortoises were collected for aerobic bacteria
isolation and submitted to the Clinical Pathology Laboratory (CPL) of the College of
Veterinary Medicine (CVM) at UF. Samples were cultured on Columbia blood agar and
MacConkeys agar, and incubated at 37C. Bacteria were identified utilizing the

32
identification systems API 20E for enteric organisms and API NFT for non-enterics
(BioMerieux Vitek, Hazelwood, MO, USA). Isolates of organisms consistent with
Pasteurella were identified to species according to biochemical profiles fisted for P.
testudinis (Snipes and Biberstein 1982).
Electron Microscopy
Selected specimens were submitted to H. P. Adams of New Mexico State
University, Las Cruces, for scanning and transmission electron microscopic preparation
and evaluation.
The left half of the bisected head of one healthy tortoise was prepared for
ultrastructural evaluations. The nasal cavity was instilled with 2.5% glutaraldehyde in 0.1
M phosphate buffer, then dissected out in its entirety, and selected areas were sampled.
The tissues were dehydrated in an ascending series of ethanols and transferred to
hexamethyldifisilazane for the final drying. The samples were sputter coated with gold and
viewed by scanning electron microscopy (SEM).
Samples from 11 diseased tortoises were collected for transmission electron
microscopy (TEM). The nasal cavity tissue was removed from the underlying
cartilaginous tissues and separated into anterior dorsal, anterior ventral, posterior dorsal
and posterior ventral quadrants. Each quadrant was cut into 1 mm cubes and placed in
2.5% glutaraldehyde, and post-fixed in osmium tetroxide. Specimens were prepared for
TEM by embedding in epon-araldite and sectioning with an ultramicrotome. Thick
sections were stained with toluidine blue and examined by fight microscopy. Ultrathin
sections were placed on copper grids, stained with uranyl acetate and lead citrate, and
examined with a Hitachi H7000 transmission electron microscope.

33
Results
Based on clinical evaluations and diagnostic tests, eight tortoises were classified as
healthy and 15 as affected by URTD. Infected tortoises were from Orange / Osceola
Counties (n = 10), Lee County (n = 3), Indian River County ( = 1) and St. Lucie County
(n = 1). Histological findings for the two groups will be discussed separately.
Normal Anatomy and Histology
The external nares opened into ventro-lateral depression, and were continuous
with large dorsal nasal cavities (Figure 3-2). Right and left dorsal nasal cavities were
separated by a cartilaginous septum. Each nasal cavity was bisected by a ridge,
forming anterior and posterior compartments. Ventrally, the nasal passageways were
continuous with the choanae (internal nares), which opened into the palatine region of
the dorsal oral cavity.
The integument continued through the external nares into a short vestibule, which
was initially lined by keratinized stratified squamous epithelium. That epithelium abruptly
changed to mucous glandular epithelium, which lined the nasal passageway throughout its
length. Interspersed among the mucous epithelial cells were ciliated epithelial cells. The
ventro-lateral depression was lined primarily by mucous and ciliated epithelial cells (Figure
3-3a). Both anterior and posterior dorsal nasal chambers were lined by a multilayered
olfactory epithelium with occasional mucous cells (Figure 3-3b). Numerous serous and
mucous glands, vessels, nerve bundles, and clusters of melanophores were present in the
connective tissue surrounding the nasal cavities. Small focal aggregates of lymphoid cells
were seen in the submucosa.

34
Figure 3-2. Diagrammatic representation of the interior of a gopher tortoise head
sectioned longitudinally, illustrating the relationship of the nasal cavity to the
external and internal nares. Approximately 3x life size. Drawing by L.
Mallory.

35
Figure 3-3. Photomicrographs of normal gopher tortoise upper respiratory tract tissues.
Hematoxylin and eosin staining, 320x. a) Photomicrograph of the ventro
lateral depression demonstrating the ciliated mucous epithelium; b)
Photomicrograph of the dorsal nasal cavity demonstrating the multilayered
olfactory epithelium. Photographs by E. R. Jacobson.

36
The bilateral thymus glands were difficult to find in healthy tortoises and were
located cranial to the base of the heart, at the branching of the subclavian and carotid
arteries. Grossly, the thymus was multilobulated. Histologically, there was a typically
dark staining cortex and a fighter staining medulla. The cortex contained densely packed
thymocytes. In the medulla there were significantly fewer cells including thymocytes, as
well as thymic epithelial cells, myoid cells, and heterophils.
The spleen was located on the right side, between the proximal duodenum and
transverse colon and was associated closely with pancreatic tissue. Histologically, spleens
were composed of distinct areas of white and red pulp. White pulp consisted of
collections of lymphoid tissue surrounding blood vessels. Red pulp, located between the
perivascular collections of the white pulp, included red blood cells within sinusoids and
small numbers of lymphocytes.
The thyroid was located at the base of the heart. In one of the healthy tortoises,
the thyroid was enlarged, with multifocal areas of follicular epithelial cell hyperplasia. In
all tortoises, follicles varied in size, with many having numerous red blood cells in the
colloid and either intraepithelial or supra-epithefial vacuoles.
Multiple foreign bodies were seen in the submucosa of the glottis of two tortoises,
in the tongue of one tortoise, and in the buccal salivary gland of one tortoise. The foreign
bodies were consistent with plant material. Lymphoid aggregates were scattered
throughout the esophagus, small intestine, large intestine and cloaca, and also were
present in the connective tissue surrounding the mental (chin) glands.
Of the healthy tortoises that were examined fully at necropsy, three were males and
three were females. In one male, multifocal areas of mineralization of seminiferous tubules

37
were seen. Golden brown granules were scattered throughout the kidney within renal
tubular epithelial cells of all tortoises. Mononuclear cells containing similar appearing
granules were within the renal interstitium and the interstitium of the testes.
Pathologic Findings
Gross examination of heads of diseased tortoises revealed minimal to large
amounts of exudate within the nasal cavity and nasal passageways. Histologically, of the
15 heads, one had no changes, two had mild changes, seven had moderate changes, and
five had severe inflammatory changes. In the tortoises with minimal changes, mild
mucosal hyperplasia and slightly increased lymphoid aggregates were seen in the nasal
passage and ventro-lateral depression. In those tortoises with moderate changes, the
olfactory epithelia were usually normal, with only focal or mild changes in the submucosa.
Changes generally were confined to the nasal passage and the ventral aspects of the nasal
cavities, and consisted of mucosal epithelial and lymphoid hyperplasia, with infiltration of
mononuclear cells and heterophils. In some tortoises with moderate inflammation, basal
cell proliferation and loss of cilia could be detected. In the tortoises with severe
inflammatory changes, there were lymphoid aggregates around submucosal glands, with
glandular epithelial hyperplasia. The normal mucosal architecture was replaced by
infiltrates of mononuclear cells and heterophils. The olfactory mucosa was replaced with
proliferating mucous epithelial cells (Figure 3-4). Proliferating basal cells projected into
the underlying lamina propria of some tortoises. Exudate, consisting of sloughed epithelial
cells and inflammatory cells, was found in the nasal cavity lumen.
Infected tortoises had larger lymphoid aggregates in the submucosa of the glottis;
several tortoises had focal areas of epithelial cell proliferation. Basal cell proliferation and

38
Figure 3-4. Photomicrograph of the nasal cavity tissues of a gopher tortoise with upper
respiratory tract disease. The changes were classified as severe, with
aggregates of lymphoid cells in the submucosa, proliferation of the basal cells,
and dysplasia of the mucosal epithelium. Hematoxylin and eosin staining,
320x. Photograph by E. R. Jacobson.

39
submucosal lymphoid hyperplasia were seen in the glottis of one tortoise. The hyperplasia
extended into the cranial tracheal epithelium. In that tortoise, there also were multifocal
areas of epithelial cell hyperplasia in the lung. Five other diseased tortoises had focal to
multifocal lymphoid aggregates in the lung interstitia.
The gastrointestinal tracts tended to have increased numbers and larger lymphoid
aggregates in the submucosa compared to those of clinically healthy tortoises. In one
tortoise there was mucous cell hyperplasia of the colon and in another there was a severe
colitis with basal epithelial cell hyperplasia and submucosal lymphoid hyperplasia, with
infiltrat es of large numbers of heterophils. Four other tortoises had increased lymphoid
aggregates in the esophagus, stomach and/or small intestine. Three additional tortoises
had increased lymphoid aggregates in the submucosa of the cloaca.
The kidneys of all diseased tortoises contained golden brown pigment granules
within renal epithelial cells. Hepatocytes of most tortoises contained similar granules, and
pathologic changes were seen in the livers of nine tortoises. In seven, there were
increased numbers and size of melanomacrophages in the liver and increased amounts of
golden brown granules. One tortoise had cuffing of the central vein by lymphocytes and
heterophils; another had aggregates of lymphocytes and melanomacrophages.
Electron micrographs of tissues of one tortoise from Sanibel Island and the tortoise
from Indian River County demonstrated organisms consistent with Mycoplasma on the
surface of the nasal mucosa (Figure 3-5). Associated epithelial cells had vacuolated
cytoplasm and inflammatory cell infiltrates were present in the mucosa. Increased
numbers of mucous epithelial cells were seen, consistent with light microscopic findings.

40
ELISA and PCR Results
ELISA and PCR results for both healthy and diseased tortoises are presented in
Table 3-1. All healthy gopher tortoises were seronegative for antibody specific against M.
agassizii. Twelve diseased tortoises were seropositive, two were suspect, and one was
seronegative. All healthy tortoises were PCR negative while 11 diseased tortoises were
PCR-positive for Mycoplasma in nasal aspirates. Four clinically healthy tortoises that had
negative culture and PCR results from nasal passage flush and swab samples had positive
cultures and / or PCR results for samples from the nasal cavities.
Microbial Isolation Results
The results of Mycoplasma and aerobic microbial cultures of the upper respiratory
tract (URT) are presented in Tables 3-1 and 3-2. Mycoplasma was not cultured from the
URT of any healthy tortoise. Mycoplasma was cultured from the URT of 11 tortoises
with URTD. The aerobic microbial isolates of healthy tortoises consisted primarily of
members of the genera Staphylococcus, Streptococcus, and Corynebacterium a few
Gram-negative rods were isolated. A greater number of Gram-negative species were
isolated from the nasal cavities of tortoises with URTD, and those isolates made up a
greater proportion of the isolates. Pasteurella testudinis was isolated from five diseased
tortoises; in two it represented the major aerobic isolate.
Discussion
Gopher tortoises with clinical signs of URTD and evidence of exposure to
Mycoplasma agassizii were obtained from multiple sites in Florida. The light microscopic

41
Figure 3-5. Transmission electron photomicrograph of the nasal cavity mucosa of a
gopher tortoise with upper respiratory tract disease. Organisms consistent
with Mycoplasma (arrow) can be seen in close association with host cell
membranes. Magnification 18,000x. Photograph by H. P. Adams.

42
Table 3-1. Summary of ELISA, PCR, culture, and nasal histopathology results from
necropsied gopher tortoises from various locations in Florida, for determining infection
with Mycoplasma agassizii.
Group
Clinical
Signs
ELISA
PCR
Mycoplasma
Culture
Histopathology
Normal
( = 8)
0%
0%
0%
0%
0%
Natural Infection
(n- 15)
60%
85%
92%
80%
92%
Table 3-2. Summary of aerobic culture results from nasal cavity swabs of gopher tortoises
from Florida.
Group
Growth
Species
Percent
of isolates
Normal
very scant to
Staphylococcus spp.
45
(n = 5)
moderate
Gram-negative rods
25
-hemolytic Streptococcus sp.
10
non-hemolytic Streptococcus spp.
10
Corynebacterium spp.
10
Natural
very scant to
Corynebacterium spp.
33
Infection
heavy
Pasteurella testudinis
18
(=11)
Staphylococcus spp.
16
Gram-negative rods
15
Micrococcus sp.
11
Flavobacterium, Pseudomonas,
7
Lactobacillus, and others

43
changes in the upper respiratory tract of the gopher tortoises were similar to inflammatory and
dysplastic changes reported for desert tortoises with URTD (Jacobson et al. 1991, Brown et
al. 1994, Jacobson et al. 1995). However, inflammationa and epithelial proliferation around
the glottis, tracheitis, and proliferative pneumonia have not been seen in desert tortoises.
Additionally, two diseased tortoises had proliferation of the colonic mucosal epithelium.
Those changes have not been seen in desert tortoises with URTD.
By electron microscopy, organisms consistent with Mycoplasma were
demonstrated on the nasal mucosal surfaces of two tortoises. Other than bacteria, no
infectious agents were demonstrated in or on nasal cavity mucosa by electron microscopy.
Eleven diseased gopher tortoises were PCR positive and M. agassizii was cultured from
the upper respiratory tract of 11 diseased tortoises examined. These results support the
hypothesis that M. agassizii is a cause of URTD in gopher tortoises in Florida.
The greater number of species and increased proportion of Gram-negative bacteria
isolated from diseased tortoises could indicate that conditions in the upper respiratory
tract of diseased tortoises are more favorable for the growth of those species, or that
tortoises infected with M. agassizii are more susceptibile to opportunistic pathogens. The
positive culture and / or PCR results from nasal cavity samples (obtained at necropsy) of
four clinically healthy tortoises with negative nasal passage flushes and swabs support the
hypothesis that tortoises can harbor the organism without showing clinical signs or
shedding bacteria. Such animals may recrudesce under stressful conditions, begin
shedding bacteria, and become infective to other tortoises.
Nine gopher tortoises exhibited pathologic changes in the livers, although the
significance of these changes is not understood currently. The mycoplasma may release

44
toxic compounds or induce the production of compounds by the tortoises that cause
damage to the liver. Desert tortoises with URTD also show changes in liver tissue (B. L.
Homer, pers. comm.), and some exhibit altered nitrogen metabolism (B. Henen, pers.
comm.). The altered nitrogen metabolism may be due to behavioral changes leading to
decreased foraging rates or efficiency, or it may be due to direct effects of the
Mycoplasma infection, but the mechanism is not yet understood. Although only one of
nine experimentally infected captive gopher tortoises significantly decreased its intake of
vegetables (G. S. McLaughlin, unpub. data), and that decrease was temporary, wild
tortoises may alter their behavior patterns to a greater degree. Alternatively, secondary
infections by other bacteria may be the proximate cause of liver damage. However, no
evidence of primary or secondary bacterial infection (i.e., necrosis) was seen histologically.
By the ELISA, none of eight clinically healthy tortoises and 11 of 15 diseased
tortoises had antibodies against M. agassizii, indicating previous exposure. Tortoises with
negative ELISA results may have been in the early stages of the disease, when increased
antibody levels had not occurred or were not detectable. The four diseased, seronegative,
tortoises may have been infected with another agent. Westhouse et al. (1996) implicated
an iridovirus as the cause of pneumonia, tracheitis, pharyngitis, and esophagitis in a gopher
tortoise from Sanibel Island. The virus was readily detectable on both light and electron
microscopy. It is not known if attempts were made to culture mycoplasma, or if an
ELISA was run on a plasma sample. No indications of viral infections were seen in the
tortoises examined for the current study.
Investigators recently have found seropositive gopher tortoises in Georgia (B.
Raphael, pers. comm.), seropositive, clinically ill, and/or PCR positive tortoises at a site in

45
northeast Florida, and seropositive tortoises at a site in Mississippi (D. M. Epperson, pers.
comm.). Although many Florida and Georgia tortoise populations are fairly large, those in
Mississippi are more restricted and are on the federal endangered species list. Outbreaks
of URTD in populations with limited recruitment and no nearby source populations could
contribute to severe declines in numbers and, possibly, local extinctions. Further studies,
on a range-wide basis, need to be conducted to determine the distribution and potential
impacts of URTD on gopher tortoise populations.
In addition, further research is necessary regarding the pathogenesis of URTD.
While findings of liver, kidney, and intestinal tract lesions are interesting, they do not
elucidate the mechanisms by which the changes are caused. Changes have been found in
hormonal profiles of some infected desert tortoises (Rostal et al. 1996), which could lead
to altered foraging and reproductive behavior, as well as decreased reproductive potential.
If foraging behavior is affected, food and water intake might be reduced, which could
affect liver and kidney functions. Both direct and indirect pathogenic mechanisms need to
be studied in order to better predict the effects of URTD on tortoises.

CHAPTER 4
EFFECTS OF REPEATED EXPOSURE ON SEROPOSITIVE ADULTS
Introduction
In order to properly evaluate the results of the diagnostic tests and incorporate
those findings into management and conservation plans, epidemiological questions,
including the response of seropositive, asymptomatic tortoises to subsequent exposure to
the agent, must be addressed. Although vaccines have been developed for some
mycoplasmal diseases (Ellison et al. 1992, Lai et al. 1996, Markham et al. 1996), they are
usually not fully protective (e.g., Djordjevic et al. 1996, Kleven et al. 1996, Mohan et al.
1996, Washburn and Weaver 1996), and many natural infections by mycoplasmas do not
engender a protective host immune response. The immune response is actually essential to
the development of lesions, and infected animals are susceptible to repeated infection
(Simecka et al. 1992). Due to the immunopathology, the disease may be more severe on
subsequent exposure than on initial infection. I designed an experiment to test the
hypothesis that tortoises that have produced antibodies against M. agassizii are protected
against reinfection with the organism and subsequent development of URTD.
46

47
Methods
Acquisition of tortoises, intake and husbandry procedures, clinical assessments,
sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed
in Chapter 2.
Statistical Analyses
The onset and severity of clinical signs of URTD, and ELISA, PCR, and culture
results for the challenge tortoises were compared to those for control and naive tortoises
using the SAS system (SAS Institute, 1988). Data from an additional naive animal
infection experiment were included after it was determined that the data did not differ
significantly from those collected from naive animals in this study. This ensured large
enough sample sizes for meaningful comparisons at more time points. Differences in the
severity of clinical signs, histologic lesions, and ELISA data among the three treatment
groups were compared by an analyses of variance-type logistic regression using maximum
likelihood estimators to compensate statistically for the different number of tortoises in
each group at each sampling date. Percentages of tortoises showing clinical signs at
different time points were compared by Fishers Exact test, with a P value of 0.05
accepted as significant.
Experimental Design
Four groups of tortoises were established. Three groups [control {n = 6), naive (n
= 11), and sentinel (n = 2)] had no history of exposure to M. agassizii, while the fourth, or
challenge group (n = 8), had previous history of exposure as indicated by a positive result

48
on the ELISA. Initially, no tortoise in any group had clinical signs of URTD, or positive
culture or PCR results. One tortoise originally slated for inclusion in the challenge group
(i.e., seropositive, but clinically, culture, and PCR negative upon arrival) developed
clinical signs before inoculation and was eliminated from the study.
Approximately 1 mo following arrival, the controls were sham inoculated
intranasally with 100 pi sterile SP4 broth in each naris, and tortoises in the naive and
challenge groups were inoculated in each naris with 100 pi of SP4 broth containing
approximately 104 colony forming units (CFU) ofM agassizii strain 723, for a total dose
of 108 CFU. The 723 isolate was obtained originally from a clinically ill tortoise from
Sanibel Island, Lee County, Florida. The sentinel tortoises were captured, but received no
other treatment.
Following inoculation, observations of all tortoises were attempted daily to
determine the onset and sequence of clinical signs. Behavior also was monitored. At 2 4
wk intervals post-inoculation (PI), tortoises were trapped, examined, and weighed, then
tranquilized and blood, nasal flush and nasal swab samples were collected. Samples were
processed as previously described. A total of 22 tortoises was examined at necropsy and
histologically. Four control and nine naive tortoises were euthanatised and necropsied in
October, 1994, before undergoing winter dormancy; seven challenge and two naive
animals were euthanatised and necropsied in March, 1995, after emergence. Complete
necropsies (n = 15) were performed on two control, six naive and seven challenge
tortoises; only heads were examined on the other animals (n 7).

49
Results
Clinical Signs
Of the six control group tortoises, none showed consistent clinical signs nor had
positive culture, PCR, or ELISA results. At each of three time points, one control
tortoise exhibited mild ocular signs that were probably associated with environmental
conditions. One of two sentinels developed clinical signs at 12 wk PI, although she had
not been inoculated with culture or medium. Of the naive tortoises, 67% exhibited clinical
signs beginning 2-3 wk PI, 79% were clinically ill by 8 wk PI, and 94% had shown signs
by 16 wk PI (Table 4-1). One naive tortoise never exhibited clinical signs. All seven
challenged tortoises developed clinical signs of URTD before dormancy. Five (71%)
challenged tortoises and one naive tortoise exhibited clinical signs soon after emergence.
No tortoises became moribund or died during the study.
At 2 wk PI, six challenge and 17 naive tortoises were examined. Challenged
tortoises had higher total clinical sign scores than naive tortoises (2.6 vs. 0.5, P < 0.001),
and higher scores for nasal discharge (1.5 vs. 0.1, P < 0.001), ocular discharge (0.8 vs. 0,
P<0.02), and palpebral edema (1.5 vs. 0.6, P < 0.02) (Table 4-1). Significantly more
challenge than naive tortoises exhibited nasal and ocular discharge (67 vs. 6%, P < 0.01;
50 vs. 0%, P < 0.02; respectively) (Figure 4-1). No consistent differences were seen
between the two groups at later sampling times (Figure 4-2).

50
Table 4-1. Percentages of naive and challenge tortoises positive for each clinical sign of
URTD, and mean clinical sign scores. Scores <1.5 are classified as slight, >1.5 and <2.5
as moderate, and >2.5 as severe. ND = nasal discharge, OD = ocular discharge, ED =
palpebral edema, CJ = conjunctivitis. P < 0.05 indicates significant difference between
groups.
Time
Sign
Naive
% Positive
Challenge
P
Naive
Mean Score
Challenge
P
2 wk
Total
67
83
0.626
0.5
2.6
0.000
NDa
6
67
0.007
0.1
1.5
0.000
OD
0
50
0.011
0
0.8
0.015
ED
59
83
0.369
0.6
1.5
0.012
CJ
53
83
0.340
0.6
1.0
0.345
4 wk
Total
79
80
1.000
1.5
1.9
0.425
ND
50
60
1.000
0.8
1.2
0.254
OD
46
20
0.366
0.8
0.6
0.666
ED
52
40
1.000
0.7
0.8
0.864
CJ
59
60
1.000
1.0
0.8
0.670
6 wk
Total
94
100
1.000
2.1
2.7
0.229
ND
67
83
0.626
1.1
1.9
0.040
OD
44
57
0.673
0.5
1.0
0.118
ED
72
71
1.000
1.0
0.9
0.652
CJ
89
43
0.032
1.4
0.7
0.055
8 wk
Total
82
83
1.000
2.6
2.4
0.688
ND
71
71
1.000
1.4
1.6
0.677
OD
54
57
1.000
0.9
0.7
0.479
ED
71
71
1.000
1.1
1.3
0.634
CJ
71
29
0.075
1.5
0.6
0.004
12 wk
Total
83
83
1.000
2.4
2.4
0.989
ND
62
71
0.642
1.5
1.6
0.836
OD
54
57
0.676
1.0
0.7
0.418
ED
67
71
0.667
0.7
1.3
0.068
CJ
79
29
0.198
1.1
0.6
0.105
Nx
Total
84
83
1.000
2.6
2.4
0.744
ND
79
71
0.646
1.4
1.6
0.733
OD
79
57
0.340
1.2
0.7
0.123
ED
58
71
0.668
1.0
1.3
0.283
CJ
63
29
0.190
1.3
0.6
0.036

90
80
70
60
50
40
30
20
10
0
4-
iNaive
CS Total ND OD
ED
CJ
Percent of gopher tortoises infected with Mycoplasma agassizii positive for any clinical sign and each individual sign at 2
weeks postinfection. Tortoises in the challenge group had serological evidence of prior exposure to M. agassizzi, but
those in the naive group did not. CS = clinical signs, ND = nasal discharge, OD = ocular discharge, ED = palpebral
edema, CJ = conjunctivitis. Indicates significant difference between groups, P < 0.05.

Mean Clinical Sign Score
Weeks Postinfection
Figure 4-2. Total clinical sign scores for three groups of tortoises, one control and two experimentally infected with Mycoplasma agassizii.
Bars represent standard errors. ** Indicates a significant (P < 0.05) rise in clinical sign score and a significant difference
between groups. Indicates a significant rise in clinical sign score.

53
Culture and PCR Results
The sentinel tortoise that became ill also had positive culture and PCR results. All
inoculated tortoises had at least two positive culture results, and 17 of 18 had positive
cultures from samples collected at necropsy. Mycoplasmal DNA was detected by PCR
analysis from each inoculated tortoise, including those that did not exhibit clinical signs of
URTD when sampled. By RFLP analysis, all isolates corresponded to M. agassizii.
ELISA Results
There was no anti-M. agassizii antibody response by the control tortoises to sham
inoculation, although one sentinel had seroconverted by 4 wk PI. All inoculated tortoises
seroconverted or had significantly increased ELISA values. ELISA values were greater
for challenge than naive tortoises at each time point (Table 4-2, Figure 4-3). A significant
increase in ELISA values of challenged tortoises was observed by 4 wk PI (mean ratio of
samples to negative control of 6.3 vs. 3.3 at 0 wk, P < 0.05), and seroconversion of naive
tortoises was observed by 6 wk PI (ratio of 3.9 vs. 1.5 at 0 wk, P < 0.05). The ratio of
the challenge to the naive ELISAs increased at 2 and 4 wk PI, then declined.
Table 4-2. Least-squares mean ELISA values for naive and challenge tortoises. P < 0.05
indicates a significant difference between groups.
Time
Naive
Challenge
P
0 weeks pi
0.2049
0.4707
0.0437
2 weeks pi
0.2235
0.5925
0.0126
4 weeks pi
0.2439
0.8848
0.0001
6 weeks pi
0.5464
1.5254
0.0001
8 weeks pi
0.8314
1.8219
0.0001
12 weeks pi
1.4062
1.9839
0.0001
Necropsy
1.6796
2.2889
0.0001

Absorbance @ 405 nm
Weeks Postinfection
Figure 4-3. ELISA results for three groups of tortoises, one control and two experimentally infected with Mycoplasma agassizii.
Tortoises in the naive group had no previous exposure to M agassizii, while those in the challenge group had evidence
of prior exposure. Line indicates ratio value (sample to negative control) of 3.0. Indicates significant differences
(P < 0.05) between challenge and naive groups. ** Indicates significant differences between naive and control groups.
*** Indicates the first significant differences from Time 0 value for that group. Bars represent standard errors.
4^

55
Histology Results
The upper respiratory tracts of the four control tortoises examined at necropsy had
normal histologic appearances. In contrast, all tortoises inoculated with mycoplasma
showed lesions similar to those seen in naturally occurring URTD. All challenge tortoises
had moderate to severe inflammation and changes in the epithelium and submucosa.
Three naive tortoises had minimal lesions and eight had moderate to severe abnormalities..
Lesions were consistently seen in the ventrolateral depression of the nasal cavity, a region
immediately caudal to the vestibule (see Chapter 3).
Discussion
No tortoises inoculated with sterile medium developed clinical signs or
histopathologic lesions, indicating that lesions were not a result of the mechanical effect of
the medium on the tissues, nor of host inflammatory response to the medium.
Experimentally infected naive animals did not seroconvert until 6 to 8 weeks PI. Because
the sentinel that became ill and seroconverted was sampled only on arrival and 8 wk
following arrival (4 wk PI for the inoculated tortoises), the timing of its seroconversion is
uncertain. No known clinically ill tortoises were transported or held with the experimental
animals once they left the Orange County holding facility, and the sentinel was not held in
the facilities or with other tortoises during the time tortoises were being inoculated.
Additionally, in an experiment involving the transfer of healthy tortoises into pens
immediately following the removal of ill tortoises, there was no indication of
environmental transfer, although the sample sizes were very small (see Chapter 7).

56
Therefore, the sentinel whose seroconversion was detected 8 wk after arrival probably
was exposed to M agassizii just before transport to UF.
Tortoises harboring M agassizii may not show clinical signs, may exhibit mild
signs, or may show signs only intermittently. Because tortoises not showing any ocular
signs or nasal discharge sometimes have positive culture and PCR results, the organism
may be transmissible from asymptomatic tortoises under appropriate conditions. Due to
unavoidable constraints on sampling live animals, I cannot exclude the possibility that
clinically healthy, seropositive, culture and PCR negative animals harbor the bacteria. As
shown by the two tortoises (the sentinel and the tortoise that was eliminated from the
study) that initially had negative culture and PCR results but developed disease without
being inoculated experimentally, some infections may go through an extended latent
period with low numbers of organisms in the nasal passages, or animals that appear to
have cleared the organism may, in fact, have not.
The clinical response of challenged animals was more rapid and severe than that of
naive animals, indicating that no protection was conferred by previous exposure to the
organism. This is consistent with some other mycoplasmal diseases in which the immune
response confers limited or no protection (Ellison et al. 1992), or contributes to
pathogenesis, such as arthritis in fowl caused by M. synoviae (Kume et al. 1977),
conjunctivitis in cattle caused by M. bovoculi (Rosenbusch 1987), and pneumonia in
humans caused by M. pneumoniae (Krause and Taylor-Robinson 1992). Tully et al.
(1995) have found that some mycoplasmal surface proteins share sequence and structural
homologies with vertebrate proteins, and suggest that these may play a role in eliciting
autoimmune responses. Repeated exposure to mycoplasmal proteins that resemble a

57
hosts proteins may sensitize the host and induce an autoimmune response, leading to
chronic manifestations of disease even if the primary etiologic agent is cleared.
Although actively produced antibodies apparently do not confer protection to adult
tortoises, it is unknown if passively transferred antibodies (see Chapter 6) provide
protection against URTD for hatchlings. Preliminary observations indicate that
seronegative hatchlings are at least as susceptible to infection as adults, and that the
disease progresses more rapidly, with high morbidity in the first 6 mo PI (see Chapter 5).
Desert tortoise hatchlings exposed to infected adults developed clinical signs of URTD
and suffered extremely high mortality rates (51 of 69, or 74%, of those with clinical signs
died) (Oftedal et al. 1996). Mycoplasma agassizii was isolated from necropsied
individuals, and severe inflammation was seen histologically. The immune status of the
hatchlings and parents was unknown prior to the development of the outbreak, so no
inferences can be drawn regarding the potential role of maternal antibodies in the course
of URTD in hatchlings. Because cellular components probably play a major role in the
pathogenesis of URTD and those are not transferred vertically in reptiles, the maternal
antibodies may provide some protection. It would be interesting to explore the question
of increased susceptibility to or severity of disease vs. protection from disease in
hatchlings that have received maternal antibodies against M. agassizii.
The immune response stimulated by previous exposure to M. agassizii did not
prevent reinfection or ameliorate signs of URTD after subsequent inoculation. The onset
of clinical signs was, in fact, more rapid and the initial severity was greater than in first
infections. Development of specific antibodies against M agassizii did not ensure
clearance of the organism by the tortoises, either on initial or subsequent exposure. Based

58
on this study, tortoises testing positive for antibodies against M agassizii cannot be
considered good candidates for release in repatriation, restocking, or relocation efforts. It
may be acceptable to release seropositive tortoises in areas that already have seropositive
animlas, as long as the overall prevalence is not increased significantly. Seropositive
tortoises may provide desirable genetic material (see Chapter 6), and would be valuable as
research subjects to study interactions with other disease agents and the effects of
repeated exposure on metabolism, autoimmune responses, reproduction, and survival.

CHAPTER 5
HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII
Introduction
In order to develop management and conservation plans that incorporate the
potential of disease to affect populations, epidemiologic questions must be addressed. Of
particular concern are the probability of transmission of the disease organism from one
tortoise to another (horizontal transmission), the rate of spread within populations, and the
potential for spread to nearby populations. The probability of transmission may be related
to a tortoises clinical and culture or PCR status. Infected tortoises may be culture and/or
PCR positive without showing clinical signs. Conversely, it is sometimes difficult to
detect bacteria in clinically ill tortoises. I designed an experiment to test the hypothesis
that horizontal transmission of M. agassizii will occur only from those individuals that are
clinically ill, and PCR and/or culture positive.
Methods
Acquisition of tortoises, intake and husbandry procedures, clinical assessments,
sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed
in Chapter 2.
59

60
Experimental Design
Fifteen pairs consisting of one male and one female were established in August,
1994. Due to the limited availability of seropositive and/or clinically ill tortoises, the
sample design was not balanced. Five pairs of asymptomatic, ELISA-, PCR-, and culture
negative control tortoises were established as controls. The other ten pairs consisted of a
clinically healthy, ELISA-, PCR-, and culture-negative tortoise that had been at UF since
April, 1994 (resident), and an ELISA-positive or -suspect (irrespective of PCR or culture
status) tortoise of the opposite sex and similar size (Figure 5-1, Table 5-1). One resident
female was paired with an ELISA-negative, but clinically ill, culture- and PCR-positive
male. The serosuspect and two seropositive tortoises were clinically healthy, and had
negative culture and PCR results. The remaining six seropositive tortoises showed
moderate clinical signs of illness. Three of those six were culture- and PCR-negative, one
was culture- and PCR-positive, one was culture-positive but PCR-negative, and one was
culture-negative and PCR-positive.
Behavioral observations and clinical signs of URTD were recorded
opportunistically (see Chapter 2), and blood and nasal flush samples were collected from
all tortoises in August and October, 1994, and in March and August, 1995. In 1996,
samples were collected in February or March, and in July or August. Clinical signs and
weights were recorded, and photographs were taken at each sampling time. Data were
analysed using Chi-square and Fisher Exact tests for differences in proportions of tortoises
becoming infected under different exposure conditions. Infected animals were
euthanatized in July, 1996, except as noted below, and uninfected tortoises were released
pursuant to Florida Game and Fresh Water Fish Commission permit number WR96224.
(

61
Figure 5-1. Flow chart showing initial distribution of seropositive, clinically ill, culture
and PCR positive tortoises.

Table 5-1. Clinical signs, culture, PCR, and ELISA status of gopher tortoises included in the upper respiratory tract disease pairing
study at each sampling time. CS clinical signs (nasal or ocular discharge, palpebral edema, and/or conjunctivitis), Cl culture results.
P PCR results, E ELISA results.
Pen
ID
Aug. 1994
Oct. 1994
Mar. 1995
Aug. 1995
Feb.-Mar. 96
Jul.-Aug. 96
Necropsy
No.
Gender
CS Cl
p
E
CS
Cl
p
E
CS
Cl
P
E
CS
Cl
P
E
CS
Cl P E
CS P
E
Date
A1
226 F
Ra
201 M
-
R
A2
577 F
4-
sb
7/96
147 M
4-c
4-
4-
-
4-
- +
4-
+
7/96
B3
261 F
-
R
160 M
R
B4
129 F
-
IT1
211 M
R
B5
108 F
R
140 M
-
R
D1
135 F
4-
4-
-
s
4-
4- +
4-
4-
7/96
241 M
-
-
-
+
-
-
-
+
-
-
-
4-
+
4-
4-
4-
+
+ +
4-
4-
7/96
D2
275 F
+
-
-
+
-
-
-
+
-
-
-
4-
+
-
-
+
-
- 4-
-
+
7/96
144 M
disappeared
If
D3
151 F
4-
+
+
+
4-
4-
s
+
+
-
4-
+
-
-
S
4-
-
- -I-
S
+
7/96
126 M
-
-
-
-
4-
+
+
4-
4-
-
4-
4-
4-
4-
-
4-
+
- 4-
-
+
7/96
D4
186 F
S
-
+
4-
-
+
+
+ +
+
+
7/96
235 M
+
+
-
+
-
+
-
4-
+
-
-
4-
-
-
+
+
-
- 4-
s
4-
7/96
D5
213 F
-
-
-
-
+
-
-
4-
+
+
+
4-
4-
4-
4-
4-
euthanatised
6/95
185 M
+
-
4-
+
+
+
+
4-
+
4-
-
4-
+
4-
-
4-
euthanatised8
6/95
D6
150 F
-
-
-
-
+
+
+
S
4-
+
+
4-
4-
S
-
4-
4-
4- 4-
+
4-
7/96
183 M
+
4-
+
-
4-
+
-
+
+
+
+
+
4-
4-
-
4-
disappeared
Uh

Table 5-1continued
Pen
ID
Aug. 1994
Oct. 1994
Mar. 1995
Aug. 1995
Feb.-Mar. 96
Jul.-Aug. 96
Necropsy
No.
Gender
CS Cl
p
E
CS
Cl
P
E
CS
Cl
P
E
CS
Cl
P E
CS
Cl
P
E
CS P
E
Date
D7
190 F
-
-
-
s
-
-
-
s
-
-
-
-
-
-
-
-
-
-
disappeared
U1
228 M
+
+
+
- +
s
-
-
+
+
+
7/96
D8
257 F
+ -
-
+
+
-
-
+
+
-
-
+
+
-
- +
+
+
-
+
+
+
7/96
130 M
-
s
+
+
s
+
+
-
+
+
+
7/96
D9
205 F
-
-
s
+
-
-
-
-
-
-
+
-
-
+
-
-
-
s
-
H
110 M
R
DIO
287 F
+ .
-
+
-
-
-
+
-
-
-
+
-
+
+
euthanatisedk
6/95
125 M
-
-
-
s
-
s
-
+
-
-
-
s
-
-
s
-
-
-
- -
-
R
aReleased pursuant to Florida Game and Fresh Water Fish Commission permit number WR96224.
bDenotes suspect status. Clinical signs: very mild, transient signs that could have been associated with environmental conditions or
mechanical irritation. Culture: presumptive colony(ies) of Mycoplasma on agar that failed to grow when transferred to broth, or were
overgrown by contamination before definite determination could be made. PCR: a very faint, nearly undetectable, signal. For ELISA,
a ratio value between 2.0 and 3.0.
cBurrowed into an adjacent pen and was exposed to an experimentally infected tortoise.
dHibemating as of December 1, 1996.
disappeared, presumed dead in burrow. Last examined August, 1995.
Euthanatised due to uterine rupture.
8Euthanatised due to poor clinical prognosis.
hDisappeared, presumed dead in burrow. Last examined March, 1996.
'Disappeared, presumed dead in burrow. Last examined May, 1996.
JUnable to evaluate ocular signs prior to administration of ketamine.
kEuthanatised due to failure to reproduce and large mass viewed on radiography.

64
Results
General Observations
Observations indicated varying degrees of interaction between pair members.
Several males were excluded from the original burrow in the pen by their female partners.
Until one of the two dug a new burrow, those males, particularly 125, 185, 201, and 235,
spent most of the day in a comer of the pen, under the shade cloth, and sometimes spent
the night out of the burrow. Other pairs spent considerable time in courtship, or foraging
in close proximity to one another.
In March, 1995, five hatchling tortoises were found in pen Dl. The female (135)
may have been gravid or storing sperm when she arrived at the UF facility in April 1994,
and laid the clutch during the summer of 1994. Alternatively, she may have copulated
with the male (241) and ovulated shortly after the pairing in August 1994. Based on
gopher tortoise physiology (Taylor 1982), however, I believe the former explanation is
more likely. One hatchling was removed and euthanatised due to congenital
abnormalities; the other four were left in the pen.
Three tortoises were euthanatised in June 1995 (Table 5-1). One, number 287
(pen DIO), was an initially seropositive, clinically ill, culture and PCR negative female
whose partner (125) had not become ill. She failed to reproduce, and a large mass could
be seen on radiographs. At necropsy, a large urolith (bladder stone), approximately 5 cm
in diameter, was removed. The second, number 213 (pen D5), was an initially
seropositive, clinically ill, culture and PCR positive female whose partner had become ill
within one month of pairing. She failed to lay six eggs, and on exploratory surgery it was
discovered that both oviducts had ruptured and the eggs were free in the coelomic cavity.

65
She was euthanatised without recovery from anesthesia. Her partner, number 185, was
clinically ill, lethargic, and losing weight, and was euthanatised shortly thereafter.
One healthy, seronegative male tortoise (144, pen D2), paired with a seropositive
female (275) disappeared after March 1995 (Table 5-1). He may have been overturned
during interactions with the female and been pushed into the burrow, where he would have
been unable to right himself. At the time of his disappearance, he had exhibited no
indications that transmission of M. agassizii had occurred. The male tortoise from pen D6
(183) also disappeared, and may have met a fate similar to that of 144. His disappearance
may have been related to agonistic interactions with any of the tortoises originally in pens
Dl, D6, D7, or D8, as discussed below. Due to the nature of tortoise burrows, it was
impossible to determine unequivocally the fates of the animals that disappeared.
Approximately 10-11 months after pairing, in June and July 1995, the burrowing
activities of the large number of tortoises in the small area resulted in the communication
of several pens (Figure 5-2). Tortoises in pens Dl, D6, D7, and D8 moved regularly
between pens and had numerous agonistic encounters with one another. The four
tortoises in pens D3 and D4 also moved between pens and had frequent interactions.
Although tortoises from pens D9 and DIO also interacted with each other, aggressive
interactions were not observed. One control male (147, pen A2), burrowed into an
adjacent pen (A7) containing an experimentally infected ill tortoise, and moved between
the two pens over several months. Neither the control female (577) nor the third tortoise
was observed to switch pens. The study was originally designed to last for 12-14 mo, but
when the animals began moving between pens, it was extended to 24 mo in order to
collect more data.

66
Figure 5-2. Burrow map showing interconnections among pens Dl, D6, D7, and D8;
between D3 and D4, and between D9 and DIO. Burrow entrances.
Evidence of Transmission of Mycoplasma agassizii
The male control tortoise (147) became ill, seroconverted, and had positive culture
and PCR results (Table 5-1). His partner became cbnically ill and seroconverted after he
returned to his original pen. No other control tortoises became ill or seroconverted. The
D-section burrowing activities mentioned above resulted in the exposure of all tortoises in
pens Dl and D7, which until that time had been clinically healthy, to clinically ill tortoises
with positive culture and/or PCR results. Three became clinically ill, and had increased
ELISA readings, with one of two initially negative tortoises seroconverting. Those
activities also resulted in the exposure of the tortoises in pen D8 to cbnically ill, culture
and PCR positive tortoises. The seronegative male (130) became ill, had positive culture
results, and seroconverted; the cbnically ill, seropositive female (257), which had been
culture and PCR negative until that time, also became culture positive. All four hatchlings
in pen Dl became severely ill, seroconverted, and had positive culture results. All were
euthanatised, and pure cultures of M. agassizii were recovered from the nasal cavities and
conjunctiva on necropsy.

67
The initially seronegative, clinically ill, culture and PCR positive male (183) had
seroconverted by October 1994. Based on his original samples from July 1994, and the
ELISA values of his October samples, he probably seroconverted within 2 wk of pairing,
and was considered as ELISA positive for the analyses (see below). In the ten
experimental pairs, eight initially asymptomatic members, two of which were originally
ELISA-positive and six of which were originally ELISA-negative, became moderately ill
with signs of URTD and had increased ELISA values, with the latter six seroconverting.
Two other tortoises (125 and 190) showed mild clinical signs that may have been
associated with mechanical irritation or environmental conditions, but did not seroconvert.
Of the four initially uninfected tortoises that showed no increase in antibody levels,
three did not have positive culture or PCR results. One (190) had only mild ocular signs,
and two (110 and 144) never showed clinical signs. The fourth (125), which consistently
showed mild signs and had a suspect PCR result in October 1994, was exposed to a
tortoise (287) that had positive samples only at necropsy. Based on observations, they
had only limited interactions. The serosuspect female (205), who was in contact with both
110 and 125, showed occasional, transient, mild clinical signs, but never had positive
cultures or PCR results, nor did her antibody levels increase. In fact at all sample times
after August 1994, she was classified as seronegative.
Of the 16 originally culture and PCR negative tortoises, 10 later had positive or
suspect culture or PCR results. Eight were exposed to clinically ill, culture or PCR
positive tortoises, seroconverted and had positive results, the ninth had positive culture
results only from necropsy samples, and the last was the individual (125) discussed above.
No tortoise became ill unless exposed to another ill tortoise.

68
Transmission Probabilities
Analysis of the data was complicated by the changing exposure status of the
tortoises due to the housing problems discussed above. Therefore, analyses were carried
out on three data sets. The first consisted of data from the first 10 mo (August 1994 -
June 1995) of the study, the second from the last 14 mo (June 1995 July 1996), and the
third of the cumulative data. The first data set (Table 5-2) consisted of the original 15
pairs. The second set (Table 5-3) did not include those pairs in which transmission had
already occurred, but only the control pairs and those whose exposure status had changed,
leaving 11 observations. The third set (Table 5-4), with 22 observations, included some
tortoises twice, as the observations of interest were the exposure events themselves, and
not the pairs or individual tortoises. Although sample sizes were very small, and some
cells had no observations, chi-squared tests of differences in proportions were run on the
three data sets. The final set was collapsed on the three predictor variables (ELISA,
clinical, and culture/PCR status; Table 5-5) to determine which had the most influence on
transmission probability.
Four exposure status classes were established based on ELISA classification,
clinical signs, and culture or PCR status. Class assignment was based on a tortoises
status at any point during the time period of interest. Therefore, a tortoises status could
change between the first and last parts of the study. Each criteria was recorded as positive
or negative, and transmission was recorded as yes or no. Transmission was classified very
conservatively, with any tortoise exhibiting even mild, transient, clinical signs not
obviously related to mechanical irritation (e.g., plant material in the eye or nose) or
environmental conditions (e.g., being held overnight in a box in which the tortoise had

69
urinated), or having one suspect culture or PCR result recorded as having been infected by
horizontal transmission. The four classes were 1) ELISA, clinically, and culture/PCR
negative (-/-/-), 2) ELISA positive, clinically and culture/PCR negative (+/-/-), 3) ELISA
and clinically positive, and culture/PCR negative (+/+/-), and 4) ELISA, clinically, and
culture/PCR positive (+/+/+). The first data set (Table 5-2) consisted of the following
observations (class, yes, no): 1,0,5; 2,0,2; 3,2,2; and 4,4,0. The second set (Table 5-3),
for the last 14 mo, was comprised of the following observations: 1,0,4; 2,0,1; 3,0,1; and
4,5,0. The last, cumulative, data set, included the following observations: 1,0,5; 2,0,3;
3,2,3; and 4,9,0. Chi-square tests of association of exposure status and transmission were
significant for all data sets, with 3 degrees of freedom (df) and P < 0.001 for all sets
(Tables 5-2, 5-3, 5-4).
Table 5-2. Contingency table for exposure status and transmission occurrence for the first
10 mo (August 1994 June 1995) of the gopher tortoise URTD pairing study.
Exposure Status
ELISA/Clinical/Culture or PCR
Transmission
Yes No
-/-/-
0 5
+1-1-
0 2
+/+/-
2 2
+/+/+
4 0
X2, 3 df= 10.8, P = 0.013, power = 0.80
Table 5-3. Contingency table for exposure status and transmission occurrence for the last
14 mo (June 1995 July 1996) of the gopher tortoise URTD pairing study.
Exposure Status
ELISA/Clinical/Culture or PCR
Transmission
Yes No
-/-/-
0 4
+1-1-
0 1
+1+1-
0 1
+1+1+
5 0
X2, 3 df = 11.0, P = 0.012, power = 0.81

70
Table 5-4. Contingency table for exposure status and transmission occurrence for the
duration (August 1994 June 1995) of the gopher tortoise URTD pairing study.
Exposure Status
ELISA/Clinical/Culture or PCR
Transmission
Yes No
-/-/-
0 5
+/-/-
0 3
+/+/-
2 3
+/+/+
9 0
X2, 3 df = 17.2, P = 0.0006, power = 0.96
Table 5-5. Transmission probabilities and tests of significant differences of proportions
for different exposure status classes of tortoises relative to upper respiratory tract disease.
Exposure
Transmission
Probability of
Fisher Exact
Status
Yes
No
Transmission
Test, P value
ELISA
negative
0
5
0
positive
11
6
0.65
0.018
Clinical illness
negative
0
8
0
positive
11
3
0.79
0.001
Culture / PCR
negative
2
11
0.15
positive
9
0
1.00
0.0002
Because transmission was defined very conservatively for the purposes of this
study, two tortoises were categorized as having been infected via horizontal transmission
based solely on the appearance of mild clinical signs. Neither animal seroconverted nor
had positive or suspect culture or PCR results during the second half of the study. Both
tortoises were housed with clinically ill, seropositive tortoises that were culture and PCR
negative. If those tortoises were designated negative relative to transmission, the final

71
data set was comprised of the following observations: 1,0,5; 2,0,3; 3,0,5; and 4,9,0. The
chi-squared value was 22.0, with 3 df and P < 0.0001.
When Table 5-4 was collapsed on ELISA status (Table 5-5), none of the tortoises
in five ELISA negative pairs became ill or seroconverted, but of the 17 interactions
involving ELISA positive tortoises, 11 resulted in transmission, for a probability of 0.65.
Fisher Exact test of a difference in proportions was significant at P 0.018. When
collapsed on clinical status, none of eight interactions involving negative tortoises resulted
in transmission, as defined above, while 11 of 14 (probability 0.79) involving clinically ill
tortoises did (Fisher exact test, P = 0.001). If transmission was defined more liberally, and
designated as positive only if a tortoise seroconverted and had clearly positive culture or
PCR results, then nine of 14 (probability 0.64) interactions with clinically ill tortoises
resulted in transmission (Fisher exact test, P < 0.006). When transmission was defined
conservatively, for culture and/or PCR status, 2 of 13 culture/PCR negative class
interactions resulted in transmission (probability 0.15), while all of nine positive class
pairings effected transmission (Fisher exact test, P = 0.0002). If transmission was defined
more liberally, then none of 13 culture/PCR negative interactions resulted in transmission,
and the Fisher exact test was significant with P = 2 x 10'6.
Discussion
The unplanned movements of the tortoises resulted in more exposure events than
initially planned, and possibly compromised some of the original experimental design.
However, at least one pair remained in each exposure category and was followed for 24

mo, twice as long as originally planned. Therefore, the intent of the study was not
compromised by the tortoises natural behavior.
Although manifestation of latent infections could not be excluded absolutely, the
results supported the hypothesis that M agassizii was horizontally transmitted between
adult tortoises, and between adult and hatchling tortoises. The route of transmission may
have been direct contact, aerosol, or fomite transmission. Direct contact was the most
likely route. If aerosol transmission occurred, it was probably over short distances, such
as would be likely to lead to direct contact. Sentinel and control animals housed in pen
groups that also housed ill tortoises did not become ill or seroconvert, indicating that
aerosolized bacteria were unlikely to travel even relatively short differences or over low
(0.5 m) barriers. Similarly, fomite transmission is unlikely to play a major role (see
Chapter 7). However, because many tortoise interactions occur in burrows, it is difficult
to assess the importance of the various modes.
Although sample sizes were small, results supported the hypothesis that
transmission is more likely when the donor is symptomatic, although tortoises without
clinical signs may be infected and able to transmit the pathogen under appropriate
conditions. After rainstorms, it is not uncommon to find two or more tortoises drinking
from the same puddle (R. E. Ashton, pers. comm.; G.S.McLaughlin, pers. obs.). When
tortoises drink, they often get water in their noses, which they then blow or sneeze out.
An asymptomatic tortoise harboring bacteria may shed enough in this manner to infect a
nearby conspecific via the aerosol route, or to leave an infective dose in the water or on
nearby plants.

73
A concurrent study (Brown et al. 1996b), in which gopher tortoises were
experimentally infected with varying doses of M. agassizii, demonstrated that the infective
dose of the 723 strain is very low, less than 10 CFU. However, an additional isolate failed
to induce disease or elicit an antibody response. In previous research involving desert
tortoises, Brown et al. (1994) demonstrated that exudate from ill tortoises elicited a
stronger immune response and induced more severe disease in inoculated tortoises than
pure cultures of M. agassizii. In addition to the mycoplasma, the exudate probably
contains other species of bacteria (e.g., Pasteurella testudinis, Coryne bacterium sp.), as
well as cellular components and cytokines that may elicit an immune response and
contribute to the disease process.
The two tortoises that exhibited mild signs without seroconverting or having
positive culture and PCR results may have been responding to cellular or chemical
components in the exudate from their partners, to environmetnal conditions, or to other
species or strains of bacteria that were only mildly pathogenic. Because conjunctival
swabs were not collected from these animals, and aerobic cultures were not performed,
those possibilities were not addressed in this study.
Further research regarding transmission probabilities and modes is needed. Larger
sample sizes and balanced designs would allow more rigorous statistical analyses.
However, establishing and maintaining such designs would be difficult, as tortoises
clinical, culture, and PCR status can vary over both the short and long term (e.g., see
Chapter 4). Experiments entailing differences in pen design could be planned to estimate
the probabilities of transmission by the various routes. For example, double layers of wire
screening between pens, with sufficient distance between the layers to prohibit direct

74
contad, could be used to assess the probability of aerosol transmission. Pen designs in
which water containers are shared, but direct contact and aerial transmission are precluded
could generate data addressing the role of transmission via water. Tortoises strength,
determination, and digging abilities must all be considered when designing such
experiments.
Seropositive, clinically healthy, culture and PCR negative individuals may be
suitable for relocation, restocking, and repatriation programs, but clinically ill animals
should not be used in such efforts. However, the proportion of tortoises that appear to
have recovered from disease and eliminated the mycoplasma, while actually harboring the
organism, is unknown. As evidenced in the preceding chapter, some of those tortoises
may have latent infections that will recrudesce under stressful conditions.

CHAPTER 6
VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII
Introduction
Many mycoplasmas can be transferred vertically, that is, from mother to offspring
in tero or in ovo (see Chapter 1). Before appropriate decisions can be made concerning
the fate of desert or gopher tortoises infected with M. agassizii, particularly those from
threatened or endangered populations, questions regarding the vertical transmissibility of
M. agassizii need to be resolved. IfM agassizii is not transmissible via eggs, the options
for preserving genetic material are increased. I designed experiments to test the
hypotheses that M. agassizii is transferred in ovo to gopher tortoise hatchlings and that
levels of specific antibodies against M. agassizii in egg yolk and hatchling plasma are
associated with those found in maternal plasma.
Methods
Adult tortoises were housed as detailed in Chapter 2, and selected as specified in
Chapter 5. In order to obtain clutches of eggs, pairs were established in August, 1994, as
detailed in Chapter 5. The sampling schedule was as detailed in Chapter 5.
75

76
Egg Collection and Incubation
Egg development was monitored by radiography (6 mas, 62-76 kv) beginning in
April of 1995 and 1996. When eggshells were judged to be calcified adequately,
oviposition was induced by intravenous injection of arginine vasotocin (Sigma) at
approximately 10-12 picograms/kg. Tortoises were monitored until clutch deposition was
complete or for a minimum of 4 hr. Eggs were collected as they were laid and placed in a
sanitized container (washed with 1:10 bleach solution and air-dried upside down) partially
filled with sterilized vermiculite moistened with an equal weight of sterilized water.
Female identification number and letter indicating order of deposition were written on
each egg with a graphite pencil swabbed with 70% ethanol. After approximately half the
clutch was laid, cloacal swabs for mycoplasma culture were taken, streaked onto SP4
agar, placed in SP4 broth, and incubated as described previously.
In 1995, eggs were incubated at 29C until hatching. In 1996, eggs were
incubated at 27, 29 or 31C. Approximately 1 wk before hatching each egg was cleansed
of adhering vermiculite with clean gauze and placed into a sanitized plastic container.
After hatching, resorption of yolk, and closing of the umbilicus, hatchlings were placed in
containers with clean sand. Hatchlings were maintained separately from adults, at ambient
temperature and light cycles, and fed natural foods supplemented with commercially
available vegetables until release into outdoor pens, where they were fed natural foods.
Culture and PCR Procedures
Eggs were taken at various times during incubation for mycoplasma culture and
antibody detection. At pipping, chorioallantoic-amniotic fluid (CAF) was collected and

77
frozen for later analyses. For culture, 100 |il samples of yolk and albumin, and small
pieces of membrane were added to 900 pi of SP4 broth. After 48 hr incubation at 30C,
500 pi of culture were removed for PCR analysis. All CAF samples obtained at pipping
were treated in the same manner. In 1996, one or two pooled samples consisting of 100
pi of CAF from each of 3 6 eggs, mixed well, were made for each clutch and processed
as described.
ELISA Procedures
Based on preliminary experiments using several methods of antibody extraction,
the supernatant resulting from mixing 1 ml of yolk with 1 ml PBS-AZ provided the most
efficient fraction for detection of antibodies (G. S. McLaughlin and I. M. Schumacher,
unpub. data). Blood samples for ELISA were collected by cardiocentesis from most
hatchlings at approximately 2-4 wk of age. In 1995, the goal was to obtain a blood
sample from each hatchling. In 1996, sampling effort was reduced to a maximum of five
samples per clutch due to the low coefficients of variation for the 1995 samples.
In 1995, hatchling ELISA samples were run as previously described except that
three dilutions from the range 1:1 to 1:8 were run in duplicate, depending on the sample
volume obtained. Sample dilutions were based on previous experiments with plasma from
desert tortoise hatchlings (I. M. Schumacher, unpub. data). All hatchling or egg samples
were run on the same plate as 1994 and 1995 samples from the corresponding female and
the presumed sire, if possible. If samples were split between two plates, plates were run
on the same day with the same reagents and positive and negative controls. In 1996,
samples were run in triplicate at a dilution of 1:2, as that was the dilution used for analysis

78
of the 1995 samples. Additionally, that modification reduced the necessary blood volume
from approximately 400 pi to 150 pi, reducing stress to the hatchlings. Samples collected
in 1995 and 1996 from the corresponding female were run on the same plate as samples
from their eggs and/or hatchlings.
Results
Clutch Sizes. Fertility and Hatching Rates
Twenty-six clutches, 13 each year, were collected. Arginine vasotocin was used to
induce opposition of 20 clutches and the remaining six clutches (five in 1995 and one in
1996) were laid in the pens. In 1995, 13 tortoises developed 115 eggs (mean = 8.8), and
103 eggs were recovered (Table 6-1). Five females laid entire clutches in the pens, four of
which were removed within 1 wk of laying. One female laid nine eggs in an inaccessible
area of her pen. Two females that laid incomplete clutches on induction laid the remaining
eggs (w 3) in their pens. Twenty-five eggs were removed for culture, PCR, and ELISA
samples. Of those, five were fertile and removed while Pable, four had died, and five
were infertile. I could not determine fertility on the remaining 11.
Overall clutch sizes (Table 6-1) were larger in 1996 (mean = 11.1) than in 1995
(mean = 8.8, 2-tail /-test = 2.78, 17 df P = 0.013), as were those of seronegative females
(11.7 vs 9.8, 2-tail /-test = 2.45, 9 df, P = 0.037). In 1996, clutch sizes of seronegative
females (mean = 11.7) were larger than those of seropositive females (mean = 10.3, 2-tail
/-test = 2.30, 11 df, P = 0.042). Thirteen females developed 144 eggs, 130 of which were
recovered. One female laid 11 eggs that were incompletely calcified and broke during the

79
laying process. Five eggs were laid in the pen and six while in transport; a usable sample
for culture and PCR was obtained from only one egg. Three females retained one egg
each, the fates of which were unknown. One fertile egg was donated for the purpose of
establishing embryonic tortoise cell lines, and 19 eggs were sampled. Of those, 13 eggs
were infertile, four had died during incubation, and I could not determine fertility of two.
In 1995, 78 eggs pipped and 77 hatchlings were produced in the laboratory (Table
6-1), and nine hatchlings were recovered from the 12 eggs laid in the pens (1/1, 1/2, 7/9).
The tortoise that pipped but did not hatch had an abnormal yolk sac; it was either
malformed or had ruptured, and yolk was found throughout the coelom. The hatchling
was in respiratory distress (cyanotic and mouth breathing), and was euthanatised. Another
fetus appeared to be full term, but had spina bifida in the cervical region, and only one
kidney.
In 1996, 111 eggs produced 113 viable hatchlings (Table 6-1). One egg produced
one healthy hatchling and one that was anencephalic. Two other eggs, from a different
clutch, each produced healthy twins. There were no differences in fertility or hatching
rates between infected and uninfected females for either year individually or both years
combined (yC tests, 1 df, all P > 0.75). Fertility and hatching rates did not differ
significantly between years (y2 tests, 1 df, all P > 0.20).
Culture and PCR Results
No cultures or PCR assays of cloacal samples, egg yolk, albumin, or membranes
were positive for mycoplasma, and no hatchlings isolated from adults developed clinical
signs (see Chapter 5 for evidence of horizontal transmission from adults to hatchlings).

80
Table 6-1. Clutch data from gopher tortoise upper respiratory tract disease vertical
transmission study, 1995 and 1996,
1995
1996
Serostatus
negative
positive
total
negative
positive
total
No. clutches
8
5
13
7
6
13
No. eggs
78
37
115
a
82
a,b
62
b
144
a
Mean
9.8
7.4
8.8
11.7
10.3
ll.l
No. recovered
75
28
103
69
61
130
No. sampled
16
9
25
11
8
19
Fertility rates
94%
97%
95%
88%
92%
90%
No. hatchlings
61
26
87
52
61
113
Hatching rates
90%
93%
91%
86%
88%
87%
CAF samples
59
19
78
51
51
102
Blood samples
51
25
76
29
28
57
aClutch sizes differed significantly between 1995 and 1996 for seronegative tortoises
(2-tail t-test = 2.45, P = 0.037), and for overall clutch size (2-tail t-test = 2.78, P =
0.013).
bClutch sizes differed significantly between seronegative and seropositive tortoises for
1996 (2-tail t-test = 2.30, P = 0.042).
ELISA Results
For ELISAs in 1995, 76 hatchlings and 34 yolks were sampled for a total of 96%
of all available eggs and hatchlings sampled. Total sampling effort per clutch ranged from
75-100%, with 50-100% of hatchlings sampled. In 1996, 57 hatchlings and 19 yolks were
sampled, for a total sampling effort of 58%, with sampling effort per clutch of42-100%.
ELISA values in egg yolks and hatchlings were correlated with maternal ELISA values in
Autumn of the prior calendar year (r = 0.68, R2 = 0.461, P < 0.0002, n = 25; Figure 6-1),
indicating that specific antibodies were transferred via the egg yolks to the hatchlings.

Hatchling antibody, OD @ 405 nm
Maternal Antibody, OD @ 405 nm
Figure 6-1. Regression of specific anti-Mycoplasma agassizii antibody levels of gopher tortoise hatchlings on maternal antibody levels,
as measured by ELISA. OD = optical density.

82
Discussion
Unlike various mycoplasmal infections of rodents and poultry (Simecka et al.
1992) and wild ducks (Goldberg et al. 1995), there was no evidence to support a
hypothesis of vertical transmission of M. agassizii. Therefore, it should be possible to
collect eggs from infected female tortoises, incubate the eggs, and release the hatchlings
with no risk of spreading the disease.
Vitellogenesis begins in August or September and continues to December (Taylor
1982), providing an extensive period for deposition of antibodies. In spite of that, the
antibody level in egg yolks and hatchling plasma was only 10 -20% of that in the maternal
plasma. Thus, there is no evidence that female gopher tortoises are sequestering
antibodies in the egg components. This is in contrast to birds, particularly chickens, which
deposit large amounts of antibodies in their eggs in a very short time period. Antibody
levels in chicken eggs can be several orders of magnitude greater than that in the maternal
plasma.
Hatchling antibody levels decline slowly over the first year of life, as the maternal
antibodies are broken down (I. M. Schumacher, unpub. data). This process is much
slower in tortoises than in mammals and birds, where such passively acquired antibodies
decline within weeks or a few months after birth or hatching. It is not known if antibodies
against M agassizii affect juvenile responses to infection with the organism Although
adult tortoises that have developed an antibody response against M. agassizii respond very
quickly and adversely to subsequent exposure (Chapter 4), that response may be mediated
more by cellular immune components (e g., macrophages and heterophils) than by

83
humoral. Hatchling and juvenile tortoises with maternal antibodies may be protected from
some effects of infection if the antibodies are neutralizing in nature, and if they can be
mobilized appropriately against the mycoplasma. Research needs to be conducted to
address these areas.

CHAPTER 7
ENVIRONMENTAL TRANSMISSION OF MYCOPLASMA AGASSIZII
Introduction
Some mycoplasmas can remain viable for at least several days in the environment
(Chandiramani et al. 1966), and for many years under refrigeration or freezing (Yoder and
Hofstad 1964). One method of controlling mycoplasma infections in domestic stock is to
depopulate a farm or facility, spray buildings and equipment with disinfectants, wait an
appropriate amount of time, and then reintroduce uninfected stock (Anonymous 1989). In
order to determine appropriate time frames for restocking tortoises, knowing the length of
time Mycoplasma agassizii remains viable in the environment is critical. Unfortunately,
direct viability testing is extremely difficult and impractical, if not impossible. Many soil
bacteria and fungi in the tortoise burrow and surrounding environment grow very rapidly
in culture and quickly overtake any mycoplasma colonies that might be present. However,
the risk of environmental transmission of M. agassizii is an important parameter in the
decision making process. In order to address this question, I designed an experiment to
test the hypothesis that environmental transmission ofM agassizii would occur in
tortoises introduced to pens previously occupied by infected, clinically ill tortoises.
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85
Methods
As a result of previous and concurrent experiments, 13 seropositive tortoises
infected with M agassizii and 15 seronegative tortoises with no clinical signs and negative
culture and PCR results were available. The 13 positive tortoises were captured and re
inoculated with approximately 108 colony forming units (CFU) ofM agassizii to ensure
active clinical disease and shedding of bacteria at the time scheduled for introducing
healthy tortoises to the pens (see Chapter 4). Six weeks following inoculation, the 13
tortoises were captured, as were the 15 healthy tortoises. Nasal flush samples were
collected from each infected tortoise to analyze by PCR.
Immediately after capture and removal of five infected tortoises from their pens, a
healthy tortoise was put into the pen. Transfers were carried out within 15 min. Transfers
were also carried out at 3 (n = 4), 7 (n = 3), and 10 (n = 3) days following removal of the
infected tortoises. Because only 13 ill tortoise pens were available, one of the day 7 and
one of the day 10 transfers consisted of pairs of tortoises. Eight weeks following transfer,
the 15 tortoises were captured, assessed clinically, photographed, and blood and nasal
flush samples were collected. Samples were handled as previously described.
Results
All infected tortoises were clinically ill at the time of capture, although PCR results
for all animals were negative. Transferred tortoises explored their new surroundings and
usually entered the burrows within 1 hr, and some entered within 15 min. At 8 wk post
transfer, no transfer tortoise was clinically ill, or had positive culture or PCR results. No
tortoise showed an increase in ELISA values indicative of possible infection (Table 7.1).

86
Table 7.1. ELISA results from initially clinically healthy, culture and PCR negative
tortoises transferred into pens previously occupied by clinically ill, culture or PCR positive
tortoises. Ratios for equal ELISA values may differ due to plate--to-plate variation in the
Tortoise
Pre-transfer ELISA
Post-transfer ELISA
Transfer
ID
Value
Ratio
Value
Ratio
Day
108
0.275
1.89
0.242
1.66
0
129
0.164
1.17
0.137
0.98
0
160
0.185
1.75
0.236
1.80
0
201
0.217
1.70
0.192
1.51
0
205
0.471
3.37
0.409
2.93
0
110
0.119
0.94
0.137
1.08
3
211
0.222
2.00
0.234
2.11
3
219
0.352
2.72
0.184
1.43
3
311
0.134
1.21
0.144
1.29
3
159
0.278
2.12
0.160
1.23
7
226
0.279
2.63
0.212
2.00
7
261
0.378
2.92
0.287
2.21
7
119
0.113
1.02
0.120
1.08
10
125
0.063
0.60
0.054
0.52
10
140
0.134
1.23
0.152
1.40
10
Discussion
Although the PCR results for the re-infected tortoises were negative at the time of
capture for transfer, the PCR is limited in its sensitivity. The lower limit of detection is
approximately 1000 CFU (D. R. Brown, unpub. data), whereas the infectious dose ofM
agassizii may be 10 or fewer CFU (Brown et al. 1996b). Therefore, tortoises could have
been shedding infective doses of bacteria and still have negative PCR results.
In other studies, 100% of 28 naive tortoises seroconverted by 8 wk PI, with all
tortoises tested at 6 wk PI having significantly increased antibody levels (Brown, D. R.,
1996b; Schumacher, unpub. data; Chapter 4). No tortoises in this study had increased

87
antibody levels, and it is unlikely that insufficient time had passed for the tortoises to
develop an immune response.
Environmental transmission of M gallisepticum is of considerable concern to the
poultry industry, necessitating disinfection of premises and equipment, and a 2-6 wk
fallow period before introducing new stock (Anonymous 1989, M. B. Brown pers.
comm.). The M. gallisepticum strain causing conjunctivitis in house finches also can
survive in the environment and cause infections in individuals later housed in the same
facility (Ley et al. 1996, Luttrell et al. 1996).
Environmental transmission of M agassizii in the wild may not be of great
concern. However, equipment used for capturing, handling, holding, and transporting
tortoises should be cleaned after each use by spraying or wiping with bleach, ethanol or
other disinfectant solution. Care should be taken to dispose properly of all gloves or
drapes that may have become contaminated. Clinically ill tortoises should not be housed
in direct contact with other animals, nor should indirect contact be allowed. Infected
tortoises should not be able to sneeze on other tortoises, nor should water or food dishes
be shared between pens without disinfection. Because the sample sizes at each time point
were very small, further research, with more rigorous attempts to isolate mycoplasmas and
quantify shedding, needs to be conducted. For the purposes of relocation, restocking, or
repatriation efforts, a short fallow period, perhaps 2 wk, is probably sufficient to prevent
environmental transmission of M. agassizii.

CHAPTER 8
CONSERVATION AND MANAGEMENT IMPLICATIONS OF
UPPER RESPIRATORY TRACT DISEASE
The research reported in this dissertation has built on and extended the findings of
research on upper respiratory tract disease in desert and gopher tortoises. Based on the
knowledge that Mycoplasma agassizii causes clinical signs and lesions of URTD and
elicits an antibody response detectable by an ELISA, experiments were designed to
compare the pathological effects of the disease in gopher tortoises to those in desert
tortoises, investigate secondary immune responses, and determine routes of transmission.
The knowledge gained can be applied to conservation and management decisions, and new
areas of research have been suggested.
Implications for Conservation and Management
When tortoises are impacted by development, mining, agriculture, or forestry
practices, decisions must be made regarding their disposition. Tortoises can be ignored,
temporarily removed and confined to pens for later return to the site, permanently moved
to a currently inhabited site, moved to a formerly occupied site, donated to research or
educational facilities, or euthanatised. The choices open for a particular population of
tortoises depend on the location, historical, current, and future site use, surrounding land
88

89
use patterns, importance of the population in maintaining genetic variability, and political
and social factors. No one prescription will be ideal for all situations, and it is difficult, if
not impossible, to develop a set of prescriptions that will cover all permutations of the
above factors. However, guidelines for making decisions can be developed, and some are
presented in this chapter.
Establishing Goals
The goals of the management action(s) must be established before decisions can be
made regarding what data to collect, or how to design survey or monitoring programs.
Management tactics designed to establish, create, or maintain a URTD-ffee population
will differ from those intended to maintain the status quo relative to disease agents. If the
genetic material represented by tortoises in defined or isolated populations (e.g., those in
South Carolina, Mississippi, or Louisiana, and some in Alabama) is important for
conservation purposes, aggressive efforts to protect and maintain the gene pool may be of
primary concern.
Understanding URTD and Test Results
An understanding of test results is necessary for proper interpretation and
application. For field personnel, recognition of clinical signs of URTD is an important
skill to gain. Differentiation of signs of URTD from other conditions is difficult even for
experts, as the signs are nonspecific, but some conditions can be recognized fairly easily
(e.g., foreign body in or trauma to the eye causing discharge, or nasal discharge due to

90
drinking or eating). Tortoise behavior interacts with clinical disease to affect transmission
probabilities, so understanding behavior also is important in the decision making process.
A clinically healthy tortoise, with negative ELISA, culture, and PCR results is
probably free from URTD. A positive ELISA, in the absence of clinical illness and
positive culture or PCR results, indicates only that the tortoise has been exposed to M.
agassizi. Because clinical signs and culture and PCR results can vary over time, we
cannot predict if or when a seropositive tortoise will break with clinical disease and begin
shedding bacteria. The more stress to which an animal is subject, such as human intrusion,
handling or transport, drought or other extreme weather conditions, the more likely it is to
have a disease recurrence. Repeated exposures to M. agassizii elicit more intense
immunological responses by the tortoises, potentially leading to autoimmune responses
that may contribute to the more severe lesions seen in longer-term infections, as well as
possible liver pathology. Tortoises with rapid responses to the mycoplasma, with copious
mucus production, may be more likely to transmit the agent to conspecifics, as their
energy reserves have not been depleted by the disease process.
A clinically ill tortoise with positive culture or PCR results, regardless of ELISA
results, probably is capable of transmitting M agassizii. The more active a tortoise is, and
the greater its daily movements, the more likely it is to spread the bacteria through a
colony and foment an outbreak of URTD. Behaviorally, male tortoises have larger home
ranges and more intraspecific contacts, so are probably at greater risk of coming in contact
with and spreading the pathogen.
V

91
A clinically healthy tortoise (i.e., one with no nasal or ocular discharge or other
signs) may have positive culture and PCR results, with either positive or negative ELISA
results. It may be in the early stages of infection or recrudescence. In the former case, the
ELISA value may be negative, but should rise within 6 wk. In the latter case, the ELISA
value may be quite high. Such tortoises may be capable of transmitting the mycoplasma
under the appropriate conditions. Although direct contact (nose-to-nose) seems to be the
most important route, transmission through water or on food cannot be ruled out. When
tortoises drink, they often expel water through their noses for short distances, up to 50
cm, or sneeze forcefully after drinking, aerosolizing the contents of the nasal passages. If
tortoises are in close contact with one another, spatially or temporally, such occurrences
may allow transmission of M. agassizii. Tortoises with slight nasal discharge, virtually
undetectable, can also aerosolize bacteria by sneezing.
Although long-term studies on the effect of URTD on survival of individuals and
populations have not been conducted, the evidence from surveys of desert tortoise
populations and from Sanibel Island indicate that the disease can have severe negative
impacts on population viability. Declines of 25-50% over 1-3 yr, and of 30-90% over 10
yr, are catastrophic for species that take 10-20 yr to reach maturity and have recruitment
rates of 1-2%. Without marked improvement in recruitment rates, affected populations
are unlikely to recover within a reasonable time frame.
Developing Questions and Conducting Surveys or Monitoring Programs
Third, questions related to management goals, and taking into account test
interpretation and tortoise behavior, must be developed. Some of the most common

92
questions are: 1) What is the prevalence of antibodies against M. agassizii in the
population of interest? 2) What is the distribution of seropositive tortoises relative to
gender, age, habitat type, geography, or land use patterns? 3) Has the seroprevalence
changed over time? 4) Is clinical disease present on the sites? 5) Is M. agassizii
detectable from any tortoises? 6) Is there evidence, either from demographic profiles or
tortoise remains, of large die-offs, or gaps in recruitment?
Once the goals have been established and questions developed, surveys or
monitoring programs can be designed to collect the necessary samples and data. After
field work has been conducted, samples collected and analysed, and test results analysed,
the information can be used to develop management programs or research questions.
When trapping tortoises for studies or relocation, it is important to note any
clinical signs that are present. Simply being aware of the possibility that URTD exists on a
site may make field personnel more likely to detect clinical cases. While this does not
guarantee that every case of URTD will be caught, it could give an early warning of
potential problems, prevent sick tortoises from being put in holding pens with healthy
ones, or transmitting the agent on equipment. It is easy to spray equipment with a mild
bleach solution, or wipe down calipers and other equipment with alcohol soaked gauze. It
is more difficult, and expensive, to construct separate holding facilities for healthy and sick
tortoises, but alternatives are available. Ill tortoises can be kept in containers for a few
days, long enough for monitoring and until test results come back.

93
Weighing Management Options and Formulating Management Plans
Knowing that URTD exists or has existed on a site, and the distribution of
seropositive tortoises, may suggest management strategies to cope with potential
problems. If tortoises testing positive are concentrated in one area, there may be
opportunities to improve habitat so tortoises are not as stressed in times of food shortage,
drought, or other adverse conditions. More monitoring time could be allocated to such
areas in order to get an early warning in the case of a disease outbreak. If the site is close
to or at carrying capacity, the population could be reduced so the remaining tortoises are
under less intraspecific stress. Eggs could be collected from that site to use in headstart
programs to ensure the genes are not lost in case of an outbreak.
If there is a population of tortoises into which relocated ones will be released, and
the goal is to maintain the status quo relative to disease agents, then both populations need
to be tested. If the recipient population and the donor population have approximately the
same level of seropositive tortoises, and the site is well below carrying capacity, then it is
probably acceptable to release seropositive tortoises, as long as the overall rate is not
increased, and no tortoises actively shedding bacteria are released.
Releasing clinically ill tortoises, regardless of ELISA results, is not recommended.
If a tortoise has a nasal discharge, palpebral edema, and conjunctivitis that persist for more
than 24 hr, there is a substantial risk that the tortoise is shedding mycoplasma. Tortoises
may shed more bacteria early in the infection, before they have developed a strong
antibody response. Due to the secondary immune responses (Chapter 4), seropositive

94
tortoises that may have cleared the bacteria would be at risk of developing severe disease
when exposed to a clinically ill tortoise, decreasing their survival probability.
If the goal is to establish, create, or maintain a URTD-ffee population, the
question remains of what to do with seropositive, clinically healthy, culture and PCR
negative individuals. If there is no secure place to hold the animals or a site where they
can be released, then the most acceptable choice may be euthanasia. Euthanasia may save
the animals from a long slow death, and prevents the chance of exposing negative animals.
Alternatively, intensive monitoring of the population, with special attention paid to
seropositive tortoises and samples collected several times per year, may allow rapid
detection and removal of ill animals. However, such careful monitoring is beyond the
capacity of most management agencies, and annual or biennial sampling may be all that
can be accomplished.
Researchers sometimes need seropositive animals for projects. If one can be
found, it is possible that animals could be donated. However, there are many issues to
consider with that approach, mostly for the researcher. Researchers usually cannot accept
animals that are made available suddenly, particularly for studies that require animals to be
kept in laboratory settings. Costs, objectives, confounding factors of length of disease,
genetics and site of origin, are all issues that must be addressed. It is expensive to keep
tortoises in captive research settings, and safeguards must be in place to prevent then-
escape or theft. Research can be a politically and socially sensitive subject due to
potentials for pain, illness, and death of animals, and nearly all animals are euthanatised at
the end of the studies.

95
If the facilities and personnel are available, and the genetic material represented by
the sick or seropositive tortoises is important, then a captive breeding program may
provide the best solution. Clinically ill animals or those that test positive by culture or
PCR should be maintained separately from clinically healthy, culture and PCR negative
animals. Eggs can be collected from the females by induction of oviposition or at natural
laying, or nests can be dug up. Eggs should not be allowed to hatch and hatchlings
emerge from nests in pens inhabited by ill adults. Hatchlings can contract URTD from
adults, and they generally become very ill very quickly. Eggs can be artificially or
naturally incubated, and hatchlings released immediately or headstarted.
Summary of Conservation and Management Implications
In summary, the following points must be considered when making conservation
and management decisions relative to URTD in gopher tortoises:
1) Goals must be clearly established.
2) Personnel must have appropriate training to recognise URTD, collect necessary
samples, and interpret results.
3) Clear questions must be formulated.
4) Survey and monitoring programs must be developed and implemented, and
precautions taken to ensure detection and prevent spread of URTD.
5) Management options must be weighed, and plans formulated and implemented
that are consistent with established goals.
a) Habitat manipulations.
b) Relocate tortoises and maintain status quo relative to URTD.

96
c) Establish, create, or maintain URTD-ffee populations by careful
monitoring, testing, and removal of infected tortoises.
d) Donate animals to research projects.
e) Establish captive breeding programs.
f) Euthanasia.
Further Research
It is not known how long Mycoplasma agassizii has been in gopher and desert
tortoises, or if it occurs naturally in tortoises in other parts of the world. At least one
other mycoplasma that causes LTRTD has been found in some populations of desert
tortoises, and there may be other etiologic agents. An iridovirus was found in an ill
gopher tortoise from Sanibel Island, and may have been the cause of the disease in that
animal (Westhouse et al. 1996). Further research needs to be conducted on that agent.
The long-term effects of M agassizii infection need to be studied. There are
indications that liver function might be affected (Chapter 3), and that reproductive
physiology is altered (Rostal et al. 1996). The impacts of altered basking and foraging
behavior need to be quantified, as do the rates of transmission within natural populations.
The role of maternal antibodies on the course of disease in hatchlings and juveniles is not
understood. In order to fully understand the impacts of URTD on tortoise populations,
many ares of research need to addressed, surveys undertaken, and long-term monitoring
programs established.

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BIOGRAPHICAL SKETCH
GRACE SHERYL McLAUGHLIN
I spent most of my life on the West Coast of North America, and attended
Humboldt State University, Areata, California, receiving a Bachelor of Arts degree in
Zoology, in spite of spending all my time in the Wildlife Department. I then ran an organic
farm for several years before moving to Ames, Iowa, to attend Iowa State University. I
studied gopher tortoises on Sanibel Island, Florida, en route to a Master of Science degree
in Animal Ecology. Before defending my thesis in November 1990,1 began my doctoral
studies at University of Florida. I originally studied parasites of bobcats and Florida
panthers, particularly their hookworms. However, I accepted a job with the gopher
tortoise upper respiratory disease project, and switched dissertation research projects. I
hope to move west and/or north soon after completion of my work on the project.
110

I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and quality,
as a dissertation for the degree of Doctor of Philosophy.
Donald J. Forrester, Chair
Professor of Wildlife Ecology and Conservation
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Yny ^ &OOW
Mary B. Brown, Cochair
Associate Professor of Veterinary Medicine
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Melvin E. Sunquist
Associate Professor of Wildlife Ecology
and Conservation
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Wiley M. Kitchens
Associate Professor of Wildlife Ecology
and Conservation
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Elliott R. Jacobson
Professor of Veterinary Medicine

I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Q. idL
Paul A. Klein
Professor of Pathology, Immunology and
Laboratory Medicine
This dissertation was submitted to the Graduate Faculty of the College of Agriculture
and to the Graduate School and was accepted as partial fulfillment of the requirements for
the degree of Doctor of Philosophy.
May 1997
guj\ X. <3*/
C2-
Dean, College of Agriculture
Dean, Graduate School



65
She was euthanatised without recovery from anesthesia. Her partner, number 185, was
clinically ill, lethargic, and losing weight, and was euthanatised shortly thereafter.
One healthy, seronegative male tortoise (144, pen D2), paired with a seropositive
female (275) disappeared after March 1995 (Table 5-1). He may have been overturned
during interactions with the female and been pushed into the burrow, where he would have
been unable to right himself. At the time of his disappearance, he had exhibited no
indications that transmission of M. agassizii had occurred. The male tortoise from pen D6
(183) also disappeared, and may have met a fate similar to that of 144. His disappearance
may have been related to agonistic interactions with any of the tortoises originally in pens
Dl, D6, D7, or D8, as discussed below. Due to the nature of tortoise burrows, it was
impossible to determine unequivocally the fates of the animals that disappeared.
Approximately 10-11 months after pairing, in June and July 1995, the burrowing
activities of the large number of tortoises in the small area resulted in the communication
of several pens (Figure 5-2). Tortoises in pens Dl, D6, D7, and D8 moved regularly
between pens and had numerous agonistic encounters with one another. The four
tortoises in pens D3 and D4 also moved between pens and had frequent interactions.
Although tortoises from pens D9 and DIO also interacted with each other, aggressive
interactions were not observed. One control male (147, pen A2), burrowed into an
adjacent pen (A7) containing an experimentally infected ill tortoise, and moved between
the two pens over several months. Neither the control female (577) nor the third tortoise
was observed to switch pens. The study was originally designed to last for 12-14 mo, but
when the animals began moving between pens, it was extended to 24 mo in order to
collect more data.


96
c) Establish, create, or maintain URTD-ffee populations by careful
monitoring, testing, and removal of infected tortoises.
d) Donate animals to research projects.
e) Establish captive breeding programs.
f) Euthanasia.
Further Research
It is not known how long Mycoplasma agassizii has been in gopher and desert
tortoises, or if it occurs naturally in tortoises in other parts of the world. At least one
other mycoplasma that causes LTRTD has been found in some populations of desert
tortoises, and there may be other etiologic agents. An iridovirus was found in an ill
gopher tortoise from Sanibel Island, and may have been the cause of the disease in that
animal (Westhouse et al. 1996). Further research needs to be conducted on that agent.
The long-term effects of M agassizii infection need to be studied. There are
indications that liver function might be affected (Chapter 3), and that reproductive
physiology is altered (Rostal et al. 1996). The impacts of altered basking and foraging
behavior need to be quantified, as do the rates of transmission within natural populations.
The role of maternal antibodies on the course of disease in hatchlings and juveniles is not
understood. In order to fully understand the impacts of URTD on tortoise populations,
many ares of research need to addressed, surveys undertaken, and long-term monitoring
programs established.


CHAPTER 8
CONSERVATION AND MANAGEMENT IMPLICATIONS OF
UPPER RESPIRATORY TRACT DISEASE
The research reported in this dissertation has built on and extended the findings of
research on upper respiratory tract disease in desert and gopher tortoises. Based on the
knowledge that Mycoplasma agassizii causes clinical signs and lesions of URTD and
elicits an antibody response detectable by an ELISA, experiments were designed to
compare the pathological effects of the disease in gopher tortoises to those in desert
tortoises, investigate secondary immune responses, and determine routes of transmission.
The knowledge gained can be applied to conservation and management decisions, and new
areas of research have been suggested.
Implications for Conservation and Management
When tortoises are impacted by development, mining, agriculture, or forestry
practices, decisions must be made regarding their disposition. Tortoises can be ignored,
temporarily removed and confined to pens for later return to the site, permanently moved
to a currently inhabited site, moved to a formerly occupied site, donated to research or
educational facilities, or euthanatised. The choices open for a particular population of
tortoises depend on the location, historical, current, and future site use, surrounding land
88


74
contad, could be used to assess the probability of aerosol transmission. Pen designs in
which water containers are shared, but direct contact and aerial transmission are precluded
could generate data addressing the role of transmission via water. Tortoises strength,
determination, and digging abilities must all be considered when designing such
experiments.
Seropositive, clinically healthy, culture and PCR negative individuals may be
suitable for relocation, restocking, and repatriation programs, but clinically ill animals
should not be used in such efforts. However, the proportion of tortoises that appear to
have recovered from disease and eliminated the mycoplasma, while actually harboring the
organism, is unknown. As evidenced in the preceding chapter, some of those tortoises
may have latent infections that will recrudesce under stressful conditions.


73
A concurrent study (Brown et al. 1996b), in which gopher tortoises were
experimentally infected with varying doses of M. agassizii, demonstrated that the infective
dose of the 723 strain is very low, less than 10 CFU. However, an additional isolate failed
to induce disease or elicit an antibody response. In previous research involving desert
tortoises, Brown et al. (1994) demonstrated that exudate from ill tortoises elicited a
stronger immune response and induced more severe disease in inoculated tortoises than
pure cultures of M. agassizii. In addition to the mycoplasma, the exudate probably
contains other species of bacteria (e.g., Pasteurella testudinis, Coryne bacterium sp.), as
well as cellular components and cytokines that may elicit an immune response and
contribute to the disease process.
The two tortoises that exhibited mild signs without seroconverting or having
positive culture and PCR results may have been responding to cellular or chemical
components in the exudate from their partners, to environmetnal conditions, or to other
species or strains of bacteria that were only mildly pathogenic. Because conjunctival
swabs were not collected from these animals, and aerobic cultures were not performed,
those possibilities were not addressed in this study.
Further research regarding transmission probabilities and modes is needed. Larger
sample sizes and balanced designs would allow more rigorous statistical analyses.
However, establishing and maintaining such designs would be difficult, as tortoises
clinical, culture, and PCR status can vary over both the short and long term (e.g., see
Chapter 4). Experiments entailing differences in pen design could be planned to estimate
the probabilities of transmission by the various routes. For example, double layers of wire
screening between pens, with sufficient distance between the layers to prohibit direct


89
use patterns, importance of the population in maintaining genetic variability, and political
and social factors. No one prescription will be ideal for all situations, and it is difficult, if
not impossible, to develop a set of prescriptions that will cover all permutations of the
above factors. However, guidelines for making decisions can be developed, and some are
presented in this chapter.
Establishing Goals
The goals of the management action(s) must be established before decisions can be
made regarding what data to collect, or how to design survey or monitoring programs.
Management tactics designed to establish, create, or maintain a URTD-ffee population
will differ from those intended to maintain the status quo relative to disease agents. If the
genetic material represented by tortoises in defined or isolated populations (e.g., those in
South Carolina, Mississippi, or Louisiana, and some in Alabama) is important for
conservation purposes, aggressive efforts to protect and maintain the gene pool may be of
primary concern.
Understanding URTD and Test Results
An understanding of test results is necessary for proper interpretation and
application. For field personnel, recognition of clinical signs of URTD is an important
skill to gain. Differentiation of signs of URTD from other conditions is difficult even for
experts, as the signs are nonspecific, but some conditions can be recognized fairly easily
(e.g., foreign body in or trauma to the eye causing discharge, or nasal discharge due to


109
Yoder, H. W., Jr. 1991. Mycoplasma galhsepticum infection, pp. 198-212 in: Calnek,
B. W., H. J. Barnes, C. W. Beard, W. M. Reid, H. W. Yoder, Jr. (eds ). Diseases
of Poultry. Iowa State University Press, Ames, IA.
Yoder, H. W., Jr., and M. S. Hofstad. 1964. Characterization of avian mycoplasma.
Avian Diseases 8: 481-512.


50
Table 4-1. Percentages of naive and challenge tortoises positive for each clinical sign of
URTD, and mean clinical sign scores. Scores <1.5 are classified as slight, >1.5 and <2.5
as moderate, and >2.5 as severe. ND = nasal discharge, OD = ocular discharge, ED =
palpebral edema, CJ = conjunctivitis. P < 0.05 indicates significant difference between
groups.
Time
Sign
Naive
% Positive
Challenge
P
Naive
Mean Score
Challenge
P
2 wk
Total
67
83
0.626
0.5
2.6
0.000
NDa
6
67
0.007
0.1
1.5
0.000
OD
0
50
0.011
0
0.8
0.015
ED
59
83
0.369
0.6
1.5
0.012
CJ
53
83
0.340
0.6
1.0
0.345
4 wk
Total
79
80
1.000
1.5
1.9
0.425
ND
50
60
1.000
0.8
1.2
0.254
OD
46
20
0.366
0.8
0.6
0.666
ED
52
40
1.000
0.7
0.8
0.864
CJ
59
60
1.000
1.0
0.8
0.670
6 wk
Total
94
100
1.000
2.1
2.7
0.229
ND
67
83
0.626
1.1
1.9
0.040
OD
44
57
0.673
0.5
1.0
0.118
ED
72
71
1.000
1.0
0.9
0.652
CJ
89
43
0.032
1.4
0.7
0.055
8 wk
Total
82
83
1.000
2.6
2.4
0.688
ND
71
71
1.000
1.4
1.6
0.677
OD
54
57
1.000
0.9
0.7
0.479
ED
71
71
1.000
1.1
1.3
0.634
CJ
71
29
0.075
1.5
0.6
0.004
12 wk
Total
83
83
1.000
2.4
2.4
0.989
ND
62
71
0.642
1.5
1.6
0.836
OD
54
57
0.676
1.0
0.7
0.418
ED
67
71
0.667
0.7
1.3
0.068
CJ
79
29
0.198
1.1
0.6
0.105
Nx
Total
84
83
1.000
2.6
2.4
0.744
ND
79
71
0.646
1.4
1.6
0.733
OD
79
57
0.340
1.2
0.7
0.123
ED
58
71
0.668
1.0
1.3
0.283
CJ
63
29
0.190
1.3
0.6
0.036


102
Howard, C. J., and R. N. Gourlay. 1978. Mycoplasmas of animals. Scientific Progress
65: 313-329.
Hudson, J. R. 1971. Contagious bovine pleuropneumonia. FAO Agricultural Studies no.
86. Food and Agricultural Organization, Rome.
Ivanics, E., R. Glavits, G. Takacs, E. Molnar, Z. Bitay, and M. Meder. 1988. An
outbreak of Mycoplasma anatis infection associated with nervous symptoms in large-
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Iverson, J. B. 1980. The reproductive biology of Gopherus polyphemus (Chelonia:
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98 in: Diemer, J. E., D. R. Jackson, J. L. Landers, J. N. Layne, and D. A. Wood,
(eds.). Gopher tortoise relocation symposium proceedings. Florida Game and Fresh
Water Fish Commission Technical Report No. 5. Tallahassee, FL.
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. Journal of Wildlife Disease 27:
296-316.
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
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Hughes. 1983. Primary late-onset hypogammaglobulinemia associated with
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respiratory disease complex. Veterinary Record 90: 556-562.


41
Figure 3-5. Transmission electron photomicrograph of the nasal cavity mucosa of a
gopher tortoise with upper respiratory tract disease. Organisms consistent
with Mycoplasma (arrow) can be seen in close association with host cell
membranes. Magnification 18,000x. Photograph by H. P. Adams.


71
data set was comprised of the following observations: 1,0,5; 2,0,3; 3,0,5; and 4,9,0. The
chi-squared value was 22.0, with 3 df and P < 0.0001.
When Table 5-4 was collapsed on ELISA status (Table 5-5), none of the tortoises
in five ELISA negative pairs became ill or seroconverted, but of the 17 interactions
involving ELISA positive tortoises, 11 resulted in transmission, for a probability of 0.65.
Fisher Exact test of a difference in proportions was significant at P 0.018. When
collapsed on clinical status, none of eight interactions involving negative tortoises resulted
in transmission, as defined above, while 11 of 14 (probability 0.79) involving clinically ill
tortoises did (Fisher exact test, P = 0.001). If transmission was defined more liberally, and
designated as positive only if a tortoise seroconverted and had clearly positive culture or
PCR results, then nine of 14 (probability 0.64) interactions with clinically ill tortoises
resulted in transmission (Fisher exact test, P < 0.006). When transmission was defined
conservatively, for culture and/or PCR status, 2 of 13 culture/PCR negative class
interactions resulted in transmission (probability 0.15), while all of nine positive class
pairings effected transmission (Fisher exact test, P = 0.0002). If transmission was defined
more liberally, then none of 13 culture/PCR negative interactions resulted in transmission,
and the Fisher exact test was significant with P = 2 x 10'6.
Discussion
The unplanned movements of the tortoises resulted in more exposure events than
initially planned, and possibly compromised some of the original experimental design.
However, at least one pair remained in each exposure category and was followed for 24


33
Results
Based on clinical evaluations and diagnostic tests, eight tortoises were classified as
healthy and 15 as affected by URTD. Infected tortoises were from Orange / Osceola
Counties (n = 10), Lee County (n = 3), Indian River County ( = 1) and St. Lucie County
(n = 1). Histological findings for the two groups will be discussed separately.
Normal Anatomy and Histology
The external nares opened into ventro-lateral depression, and were continuous
with large dorsal nasal cavities (Figure 3-2). Right and left dorsal nasal cavities were
separated by a cartilaginous septum. Each nasal cavity was bisected by a ridge,
forming anterior and posterior compartments. Ventrally, the nasal passageways were
continuous with the choanae (internal nares), which opened into the palatine region of
the dorsal oral cavity.
The integument continued through the external nares into a short vestibule, which
was initially lined by keratinized stratified squamous epithelium. That epithelium abruptly
changed to mucous glandular epithelium, which lined the nasal passageway throughout its
length. Interspersed among the mucous epithelial cells were ciliated epithelial cells. The
ventro-lateral depression was lined primarily by mucous and ciliated epithelial cells (Figure
3-3a). Both anterior and posterior dorsal nasal chambers were lined by a multilayered
olfactory epithelium with occasional mucous cells (Figure 3-3b). Numerous serous and
mucous glands, vessels, nerve bundles, and clusters of melanophores were present in the
connective tissue surrounding the nasal cavities. Small focal aggregates of lymphoid cells
were seen in the submucosa.


85
Methods
As a result of previous and concurrent experiments, 13 seropositive tortoises
infected with M agassizii and 15 seronegative tortoises with no clinical signs and negative
culture and PCR results were available. The 13 positive tortoises were captured and re
inoculated with approximately 108 colony forming units (CFU) ofM agassizii to ensure
active clinical disease and shedding of bacteria at the time scheduled for introducing
healthy tortoises to the pens (see Chapter 4). Six weeks following inoculation, the 13
tortoises were captured, as were the 15 healthy tortoises. Nasal flush samples were
collected from each infected tortoise to analyze by PCR.
Immediately after capture and removal of five infected tortoises from their pens, a
healthy tortoise was put into the pen. Transfers were carried out within 15 min. Transfers
were also carried out at 3 (n = 4), 7 (n = 3), and 10 (n = 3) days following removal of the
infected tortoises. Because only 13 ill tortoise pens were available, one of the day 7 and
one of the day 10 transfers consisted of pairs of tortoises. Eight weeks following transfer,
the 15 tortoises were captured, assessed clinically, photographed, and blood and nasal
flush samples were collected. Samples were handled as previously described.
Results
All infected tortoises were clinically ill at the time of capture, although PCR results
for all animals were negative. Transferred tortoises explored their new surroundings and
usually entered the burrows within 1 hr, and some entered within 15 min. At 8 wk post
transfer, no transfer tortoise was clinically ill, or had positive culture or PCR results. No
tortoise showed an increase in ELISA values indicative of possible infection (Table 7.1).


93
Weighing Management Options and Formulating Management Plans
Knowing that URTD exists or has existed on a site, and the distribution of
seropositive tortoises, may suggest management strategies to cope with potential
problems. If tortoises testing positive are concentrated in one area, there may be
opportunities to improve habitat so tortoises are not as stressed in times of food shortage,
drought, or other adverse conditions. More monitoring time could be allocated to such
areas in order to get an early warning in the case of a disease outbreak. If the site is close
to or at carrying capacity, the population could be reduced so the remaining tortoises are
under less intraspecific stress. Eggs could be collected from that site to use in headstart
programs to ensure the genes are not lost in case of an outbreak.
If there is a population of tortoises into which relocated ones will be released, and
the goal is to maintain the status quo relative to disease agents, then both populations need
to be tested. If the recipient population and the donor population have approximately the
same level of seropositive tortoises, and the site is well below carrying capacity, then it is
probably acceptable to release seropositive tortoises, as long as the overall rate is not
increased, and no tortoises actively shedding bacteria are released.
Releasing clinically ill tortoises, regardless of ELISA results, is not recommended.
If a tortoise has a nasal discharge, palpebral edema, and conjunctivitis that persist for more
than 24 hr, there is a substantial risk that the tortoise is shedding mycoplasma. Tortoises
may shed more bacteria early in the infection, before they have developed a strong
antibody response. Due to the secondary immune responses (Chapter 4), seropositive


36
The bilateral thymus glands were difficult to find in healthy tortoises and were
located cranial to the base of the heart, at the branching of the subclavian and carotid
arteries. Grossly, the thymus was multilobulated. Histologically, there was a typically
dark staining cortex and a fighter staining medulla. The cortex contained densely packed
thymocytes. In the medulla there were significantly fewer cells including thymocytes, as
well as thymic epithelial cells, myoid cells, and heterophils.
The spleen was located on the right side, between the proximal duodenum and
transverse colon and was associated closely with pancreatic tissue. Histologically, spleens
were composed of distinct areas of white and red pulp. White pulp consisted of
collections of lymphoid tissue surrounding blood vessels. Red pulp, located between the
perivascular collections of the white pulp, included red blood cells within sinusoids and
small numbers of lymphocytes.
The thyroid was located at the base of the heart. In one of the healthy tortoises,
the thyroid was enlarged, with multifocal areas of follicular epithelial cell hyperplasia. In
all tortoises, follicles varied in size, with many having numerous red blood cells in the
colloid and either intraepithelial or supra-epithefial vacuoles.
Multiple foreign bodies were seen in the submucosa of the glottis of two tortoises,
in the tongue of one tortoise, and in the buccal salivary gland of one tortoise. The foreign
bodies were consistent with plant material. Lymphoid aggregates were scattered
throughout the esophagus, small intestine, large intestine and cloaca, and also were
present in the connective tissue surrounding the mental (chin) glands.
Of the healthy tortoises that were examined fully at necropsy, three were males and
three were females. In one male, multifocal areas of mineralization of seminiferous tubules


90
drinking or eating). Tortoise behavior interacts with clinical disease to affect transmission
probabilities, so understanding behavior also is important in the decision making process.
A clinically healthy tortoise, with negative ELISA, culture, and PCR results is
probably free from URTD. A positive ELISA, in the absence of clinical illness and
positive culture or PCR results, indicates only that the tortoise has been exposed to M.
agassizi. Because clinical signs and culture and PCR results can vary over time, we
cannot predict if or when a seropositive tortoise will break with clinical disease and begin
shedding bacteria. The more stress to which an animal is subject, such as human intrusion,
handling or transport, drought or other extreme weather conditions, the more likely it is to
have a disease recurrence. Repeated exposures to M. agassizii elicit more intense
immunological responses by the tortoises, potentially leading to autoimmune responses
that may contribute to the more severe lesions seen in longer-term infections, as well as
possible liver pathology. Tortoises with rapid responses to the mycoplasma, with copious
mucus production, may be more likely to transmit the agent to conspecifics, as their
energy reserves have not been depleted by the disease process.
A clinically ill tortoise with positive culture or PCR results, regardless of ELISA
results, probably is capable of transmitting M agassizii. The more active a tortoise is, and
the greater its daily movements, the more likely it is to spread the bacteria through a
colony and foment an outbreak of URTD. Behaviorally, male tortoises have larger home
ranges and more intraspecific contacts, so are probably at greater risk of coming in contact
with and spreading the pathogen.
V


5
tortoises was a major impact on regional distribution, and current poaching activities may
extirpate local aggregations (Diemer 1989, Mann 1990). Large scale conversion of long-
leaf pine habitats to slash pine plantations, transformation of native pasture or savanna to
improved pasture, citrus groves, or row crops, mining operations such as phosphate,
mineral sand, and gravel mines, and urban/suburban development are the main threats to
continued gopher tortoise survival today (Diemer 1986, Cox et al. 1987, Diemer and
Moore 1994). Lack of natural fire regimes due to suppression efforts by humans may alter
vegetation mosaics in remaining habitat, rendering them less suitable to continued
maintenance of tortoise populations (Mushinsky 1986, Mushinsky and Gibson 1991).
Although tortoise numbers increase with increasing areal extent of available
habitat, densities remain constant or decrease (Mushinsky and McCoy 1994).
Fragmentation of mainland areas, with ever smaller islands of suitable habitat surrounded
by agricultural and urban development, may force tortoises into higher density populations
than would occur normally. Vegetational changes resulting from reduced fire incidence in
small areas, particularly increased canopy closure and decreased herbaceous vegetation
(Mushinsky 1985, 1986), can lead to decreases in reproductive rates and juvenile survival
(Aufifenberg and Franz 1982, McLaughlin 1990, Mushinsky and McCoy 1994). Increased
intraspecific interactions at higher densities (McRae et al. 1981) may lead to elevated
physiological stress, potentially affecting immune system function and rendering animals
more susceptible to disease. Increased densities and number of interactions multiply the
opportunities for transmission of communicable agents, making fragmented populations
more likely to sustain high morbidity during epizootics. If mortality is high, lowered
reproduction due to habitat degradation may be insufficient for population recovery.


99
Cassell, G. H., J. K. Davis, N. R. Cox, M. K. Davidson, and J. R. Lindsey. 1984.
Mycoplasma pulmonis detection in rodents: lessons for diagnosis in other species.
Israel Journal of Medical Science 20: 859-865.
Chandiramani, N. K., H. Van Roekel, and O. M. Olesiuk. 1966. Viability studies with
Mycoplasma eallisepticum under different environmental conditions. Poultry Science
45: 1029-1044.
Chanock, R. M., L. Dienes, M. D. Eaton, D. G. Edward, E. A. Freundt, L. Hayflick, J. F.
P. Hers, K. E. Jensen, C. Liu, B. P. Marmion, H. E. Morton, M. A. Mufson, P. F.
Smith, N. L. Somerson, and D. Taylor-Robinson. 1963. Mycoplasma pneumoniae:
proposed nomenclature for atypical pneumonia organism (Eatons agent). Science
140: 662.
Clyde, W. A. Jr. 1979. Mycoplasma pneumoniae infections of man. Pp. 275-306 in J. G.
Tully and R. F. Whitcomb (eds.). The Mycoplasmas, vol 2. Academic Press, New
York.
Clyde, W. A. Jr. 1983. Mycoplasma pneumoniae respiratory disease symposium:
summation and significance. Yale Journal of Biological Medicine 56: 523-527.
Com, P. S. 1994. Recent trends of desert tortoise populations in the Mojave Desert. Pp.
85-94 in R. B. Bury and D. J. Germano (eds.). Biology of North American tortoises.
National Biological Survey, Fish and Wildlife Research Report 13.
Cottew, G. S. 1984. Overview of mycoplasmoses in sheep and goats. Israel Journal of
Medical Science 20: 962-964.
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. Florida Game and Fresh Water Fish Commission Nongame
Wildlife Program, Technical Report No. 4. Tallahassee, FL. 75 pp.
Dajani, A. S., W. A. Clyde, Jr., and F. W. Denny. 1965. Experimental infection with
Mycoplasma pneumoniae (Eatons agent). Journal of Experimental Medicine 121:
1071-1084.
Davidson, W. R., V. F. Nettles, C. E. Couvillion, and H. W. Yoder. 1982. Infectious
sinusitis in wild turkeys. Avian Diseases 26: 402-405.
Davidson, W. R., H. W. Yoder, M. Brugh, and V. F. Nettles. 1988. Serological
monitoring of Eastern Wild Turkeys for antibodies to Mycoplasma spp. and avian
influenza viruses. Journal of Wildlife Diseases 24: 348-351.


mo, twice as long as originally planned. Therefore, the intent of the study was not
compromised by the tortoises natural behavior.
Although manifestation of latent infections could not be excluded absolutely, the
results supported the hypothesis that M agassizii was horizontally transmitted between
adult tortoises, and between adult and hatchling tortoises. The route of transmission may
have been direct contact, aerosol, or fomite transmission. Direct contact was the most
likely route. If aerosol transmission occurred, it was probably over short distances, such
as would be likely to lead to direct contact. Sentinel and control animals housed in pen
groups that also housed ill tortoises did not become ill or seroconvert, indicating that
aerosolized bacteria were unlikely to travel even relatively short differences or over low
(0.5 m) barriers. Similarly, fomite transmission is unlikely to play a major role (see
Chapter 7). However, because many tortoise interactions occur in burrows, it is difficult
to assess the importance of the various modes.
Although sample sizes were small, results supported the hypothesis that
transmission is more likely when the donor is symptomatic, although tortoises without
clinical signs may be infected and able to transmit the pathogen under appropriate
conditions. After rainstorms, it is not uncommon to find two or more tortoises drinking
from the same puddle (R. E. Ashton, pers. comm.; G.S.McLaughlin, pers. obs.). When
tortoises drink, they often get water in their noses, which they then blow or sneeze out.
An asymptomatic tortoise harboring bacteria may shed enough in this manner to infect a
nearby conspecific via the aerosol route, or to leave an infective dose in the water or on
nearby plants.


Table 5-1continued
Pen
ID
Aug. 1994
Oct. 1994
Mar. 1995
Aug. 1995
Feb.-Mar. 96
Jul.-Aug. 96
Necropsy
No.
Gender
CS Cl
p
E
CS
Cl
P
E
CS
Cl
P
E
CS
Cl
P E
CS
Cl
P
E
CS P
E
Date
D7
190 F
-
-
-
s
-
-
-
s
-
-
-
-
-
-
-
-
-
-
disappeared
U1
228 M
+
+
+
- +
s
-
-
+
+
+
7/96
D8
257 F
+ -
-
+
+
-
-
+
+
-
-
+
+
-
- +
+
+
-
+
+
+
7/96
130 M
-
s
+
+
s
+
+
-
+
+
+
7/96
D9
205 F
-
-
s
+
-
-
-
-
-
-
+
-
-
+
-
-
-
s
-
H
110 M
R
DIO
287 F
+ .
-
+
-
-
-
+
-
-
-
+
-
+
+
euthanatisedk
6/95
125 M
-
-
-
s
-
s
-
+
-
-
-
s
-
-
s
-
-
-
- -
-
R
aReleased pursuant to Florida Game and Fresh Water Fish Commission permit number WR96224.
bDenotes suspect status. Clinical signs: very mild, transient signs that could have been associated with environmental conditions or
mechanical irritation. Culture: presumptive colony(ies) of Mycoplasma on agar that failed to grow when transferred to broth, or were
overgrown by contamination before definite determination could be made. PCR: a very faint, nearly undetectable, signal. For ELISA,
a ratio value between 2.0 and 3.0.
cBurrowed into an adjacent pen and was exposed to an experimentally infected tortoise.
dHibemating as of December 1, 1996.
disappeared, presumed dead in burrow. Last examined August, 1995.
Euthanatised due to uterine rupture.
8Euthanatised due to poor clinical prognosis.
hDisappeared, presumed dead in burrow. Last examined March, 1996.
'Disappeared, presumed dead in burrow. Last examined May, 1996.
JUnable to evaluate ocular signs prior to administration of ketamine.
kEuthanatised due to failure to reproduce and large mass viewed on radiography.


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4 EFFECTS OF REPEATED EXPOSURE ON
SEROPOSITIVE ADULTS 46
Introduction 46
Methods 47
Statistical Analyses 47
Experimental Design 47
Results 49
Clinical Signs 49
Culture and PCR Results 53
ELISA Results 53
Histology Results 55
Discussion 55
5 HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII . 59
Introduction 59
Methods 59
Experimental Design 60
Results 64
General Observations 64
Evidence of Transmission of Mycoplasma agassizii 66
Transmission Probabilities 68
Discussion 71
6 VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII 75
Introduction 75
Methods 75
Egg Collection and Incubation 76
Culture and PCR Procedures 76
ELISA Procedures 77
Results 78
Clutch Sizes, Fertibty and Hatching Rates 78
Culture and PCR Results 79
ELISA Results 80
Discussion 82
7 ENVIRONMENTAL TRANSMISSION OF
MYCOPLASMA AGASSIZII 84
Introduction 84
Methods 84
Results 85
Discussion 86
vi


105
Markham, J. F., P. C. Scott, and K. G. Whithear. 1996. Field and laboratory studies on a
live attenuated Mycoplasma synoviae vaccine. IOM Letters 4: 286.
McLaughlin, G. S. 1990. Ecology of gopher tortoises (Gopherus polyphemus) on
Sanibel Island, Florida. M. S. Thesis. Iowa State Univ., Ames. 124 pp.
McRae, W. A., J. L. Landers, and J. A. Gamer. 1981. Movement patterns and home
range of the gopher tortoise. American Midland Naturalist 106: 165-179.
Mohan, K., C. M. Foggin, P. Muvavarirwa, J. Honywill, and A. Pawandiwa. 1995.
Mycoplasma-associated polyarthritis in farmed crocodiles (Crocodylus niloticus) in
Zimbabwe. Onderstepoort Journal of Veterinary Research 62:45-49.
Mohan, K., C. M. Foggin, P. Muvavarirwa, and J. Honywill. 1996. Experimental trial
with an alum-precipitated vaccine against Mycoplasma-associated polyarthritis in
farmed crocodiles (Crocodylus niloticus). IOM Letters 4: 287.
Murray, H. W., H. Masur, L. B. Senterfit, and R. B. Roberts. 1975. The protean
manifestations of Mycoplasma pneumoniae infections in adults. American Journal of
Medicine 58: 229-242.
Mushinsky, H. R. 1985. Fire and the Florida sandhill herpetofaunal community: with
special attention to responses of Cnemidophorous sexlineatus. Herpetologica 41:
333-342.
Mushinsky, H. R. 1986. Fire, vegetation structure, and herpetofaunal communities. Pp.
383-388 in Z. Rocek (ed.). Studies in herpetology. Charles University Press, Prague.
Mushinsky, H. R, and D. J. Gibson. 1991. The influence of fire on habitat structure.
Pp. 237-259 in S. S. Bell, E. D. McCoy, and H. R. Mushinsky (eds.). The physical
arrangement of objects in space. Chapman and Hall, London.
Mushinsky, H. R, and E. D. McCoy. 1994. Comparison of gopher tortoise populations on
islands and on the mainland in Florida. Pp. 113-128 in R. B. Bury and D. J. Germano
(eds.). Biology of North American tortoises. National Biological Survey, Fish and
Wildlife Research 13.
Mushinsky, H. R, D. S. Wilson, and E. D. McCoy. 1994. Growth and sexual
dimorphism of Gopherus polyphemus in central Florida. Herpetologica 50: 119-128.
Naftalin, J. M., G. Wellish, Z. Kanana, and D. Diengott. 1974. Mycoplasma pneumoniae
septicemia. Journal of the American Medical Association 228: 565.
Nettles, V. 1996. Reemerging and emerging infectious diseases: Economic and other
impacts on wildlife. ASM News 62: 589-591.


CHAPTER 3
NATURALLY OCCURRING UPPER RESPIRATORY TRACT DISEASE
Methods
Tortoises
Twenty-three gopher tortoises from the following locations in Florida (Figure 3-1)
were transported to UF from August 1993 to September 1995: Alachua County (n = 2),
Sanibel Island, Lee County (n = 3), Volusia County (n = 1), St. Lucie County (n= 1),
Indian River County (n = 1), Orange and/or Osceola Counties (n = 15). Collection of
tortoises, except those from Orange and Osceola Counties, was opportunistic, and a result
of submissions to the UF Wildlife Clinic, or other veterinary clinics. Some tortoises had
exhibited signs of URTD, while others had been hit by automobiles. Tortoises from
Orange and Osceola Counties were selected on the basis of clinical evaluations, ELISA,
culture, and / or PCR results. Six clinically healthy animals were included in the latter
group. Tortoises were evaluated for clinical signs of URTD (i.e., nasal and ocular
discharge, palpebral edema, and conjunctivitis) and those exhibiting one or more signs of
disease, with a past history of clinical signs, with positive culture or PCR results, or with a
positive ELISA result, were designated as diseased. Tortoises were designated healthy if
they were free of any history of or current signs of URTD and were culture, PCR, and
ELISA negative.
30


31
Figure 3-1. Locations in Florida from which gopher tortoises were obtained. 1 Alachua
County, 2 Volusia County, 3 Orange/Osceola Counties, 4 Indian River
County, 5 St. Lucie County, 6 Sanibel Island, Lee County.
Necropsy and Histology Procedures
Necropsy procedure and light microscopic evaluation of tissues were performed as
detailed in Chapter 2. Gross necropsies were conducted on 21 tortoises. Multiple tissues
were collected from 15 tortoises, heads and livers from six and only heads from two.
Microbial Isolation
Flush and swab samples for Mycoplasma isolation were collected from the nasal
passages and cavities of each tortoise and processed as described in Chapter 2. Swab
specimens of the dorsal nasal cavities of 16 tortoises were collected for aerobic bacteria
isolation and submitted to the Clinical Pathology Laboratory (CPL) of the College of
Veterinary Medicine (CVM) at UF. Samples were cultured on Columbia blood agar and
MacConkeys agar, and incubated at 37C. Bacteria were identified utilizing the


20
infected host that is clinically ill and PCR or culture positive than from a seropositive
host that is not clinically ill or is clinically ill but culture or PCR negative (Chapter 5).
4) To determine ifM agassizii is transmitted vertically (Chapter 6).
5) To assess the relationship of antibodies in eggs and hatchling serum to those in
maternal serum (Chapter 6).
6) To collect preliminary data to test the hypothesis that M. agassizii can be transmitted
environmentally (e.g., in burrows) (Chapter 7).
7) To discuss the implications of the findings of the above research and additional
concurrent research on conservation and management of gopher and other tortoises
(Chapter 8).


28
concentrated barbiturate solution (Socumb, The Butler Company, Columbus, Ohio, USA)
intraperitoneally at 1 ml/kg. Once the tortoises showed complete muscle relaxation and
were unresponsive to painful stimulation, they were exsanguinated via a 23 gauge butterfly
catheter inserted into the carotid artery and then decapitated. Flush and swab samples
were collected as previously described, then the head was bisected longitudinally with an
electric saw. Following bisection, the cartilage over each nasal cavity was reflected
aseptically, and flushes and swabs of 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 and the following tissues were collected, fixed in neutral buffered 10%
formalin, sectioned at 5-6 pm, and stained with hematoxylin and eosin: glottis, cranial
trachea, tracheal bifurcation, left lung, right lung, thyroid, heart, brain, thymus, esophagus,
stomach, small intestine, pancreas, large intestine, cloaca, spleen, liver, left and right
kidney, bladder, right and left gonads, chin gland, buccal salivary gland, chin gland, and
tongue. Tissues were examined by light microscopy and abnormalities or changes were
recorded.
Histopathology Procedures
For histopathologic studies, heads were fixed in 10% neutral buffered formalin
(NBF), decalcified, embedded in paraffin, sectioned longitudinally at 5-6 pm, and stained
with hematoxylin and eosin. Sections were examined by light microscopy and classified on
a scale of 0 to 5, with 0 being normal and 5 exhibiting severe inflammation and / or
changes. Changes in the epithelium and submucosa were recorded separately.


48
on the ELISA. Initially, no tortoise in any group had clinical signs of URTD, or positive
culture or PCR results. One tortoise originally slated for inclusion in the challenge group
(i.e., seropositive, but clinically, culture, and PCR negative upon arrival) developed
clinical signs before inoculation and was eliminated from the study.
Approximately 1 mo following arrival, the controls were sham inoculated
intranasally with 100 pi sterile SP4 broth in each naris, and tortoises in the naive and
challenge groups were inoculated in each naris with 100 pi of SP4 broth containing
approximately 104 colony forming units (CFU) ofM agassizii strain 723, for a total dose
of 108 CFU. The 723 isolate was obtained originally from a clinically ill tortoise from
Sanibel Island, Lee County, Florida. The sentinel tortoises were captured, but received no
other treatment.
Following inoculation, observations of all tortoises were attempted daily to
determine the onset and sequence of clinical signs. Behavior also was monitored. At 2 4
wk intervals post-inoculation (PI), tortoises were trapped, examined, and weighed, then
tranquilized and blood, nasal flush and nasal swab samples were collected. Samples were
processed as previously described. A total of 22 tortoises was examined at necropsy and
histologically. Four control and nine naive tortoises were euthanatised and necropsied in
October, 1994, before undergoing winter dormancy; seven challenge and two naive
animals were euthanatised and necropsied in March, 1995, after emergence. Complete
necropsies (n = 15) were performed on two control, six naive and seven challenge
tortoises; only heads were examined on the other animals (n 7).


55
Histology Results
The upper respiratory tracts of the four control tortoises examined at necropsy had
normal histologic appearances. In contrast, all tortoises inoculated with mycoplasma
showed lesions similar to those seen in naturally occurring URTD. All challenge tortoises
had moderate to severe inflammation and changes in the epithelium and submucosa.
Three naive tortoises had minimal lesions and eight had moderate to severe abnormalities..
Lesions were consistently seen in the ventrolateral depression of the nasal cavity, a region
immediately caudal to the vestibule (see Chapter 3).
Discussion
No tortoises inoculated with sterile medium developed clinical signs or
histopathologic lesions, indicating that lesions were not a result of the mechanical effect of
the medium on the tissues, nor of host inflammatory response to the medium.
Experimentally infected naive animals did not seroconvert until 6 to 8 weeks PI. Because
the sentinel that became ill and seroconverted was sampled only on arrival and 8 wk
following arrival (4 wk PI for the inoculated tortoises), the timing of its seroconversion is
uncertain. No known clinically ill tortoises were transported or held with the experimental
animals once they left the Orange County holding facility, and the sentinel was not held in
the facilities or with other tortoises during the time tortoises were being inoculated.
Additionally, in an experiment involving the transfer of healthy tortoises into pens
immediately following the removal of ill tortoises, there was no indication of
environmental transfer, although the sample sizes were very small (see Chapter 7).


knew and taught me much. Many of the VMTH personnel, especially Neil, Danielle, Tom,
Dana, and An have assisted and taught me, and cheered me up when I needed it.
Ms. Joan Berish has provided an incredible amount of advice, support, and
friendship throughout the eight years I have been studying gopher tortoises, and I only
hope to reciprocate. Garry Fosters assistance through my early years at UF were
invaluable, as were the support of my fellow graduate students in Dr. Forresters lab,
Marisol Sepulveda and Don Coyner. My brother Mark, my sister Megan, and friends
Vicky, Bill, Sharon, Kay, Pierre, Andrea, and Tania helped me get here and stick it out,
and deserve thanks for their support. There have been several other people who have
helped me at various times and in different ways, especially those of the gopher tortoise
sodality, and my thanks go to them also. I also thank the members and friends of the
Unitarian Universalist Fellowship of Gainesville for being my family and caring for me.
I was supported by a Presidential Research Fellowship and a Gatorade Grant from
the University of Florida, and funds from The Walt Disney World Company.
IV


100
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States. Herpetologica 42:125-133.
Diemer, J. E. 1989. Gopher tortoise status and harvest impact determination. Final
report. Florida Game and Fresh Water Fish Commission, Tallahassee, FL. 320 pp.
Diemer, J. E. 1992a. Gopher tortoise, pp. 123-127 in: Moler, P. E. (ed.) Rare and
endangered biota of Florida volume III: amphibians and reptiles. University of Florida
Press, Gainesville, FL.
Diemer, J. E. 1992b. Home range and movements of the tortoise Gopherus polyphemus
in northern Florida. Herpetologica 42: 158-165.
Diemer, J. E., and C. L. Moore. 1994. Reproduction of gopher tortoises in north-central
Florida. Pp. 129-137 in R. B. Bury and D. J. Germano (eds.). Biology of North
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Dietlein, N. E., and R. Franz. 1979. Status and habits of Gopherus polyphemus. Pp.
175-180 in E. St. Amant (ed.). Desert Tortoise Council Proceedings 1979
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Djordjevic, S. P., G. J. Eamens, A. L. Scarman, and J. C. Chin. 1996. Characterisation of
Mycoplasma hyopneumoniae membrane proteins and identification of discrete
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a virulent strain ofM hyopneumoniae. IOM Letters 4: 282-283.
do Nascimento, M. da G., and E. R. do Nascimento. 1986. Infectious sinusitis in
cotumix quail in Brazil. Avian Diseases 30: 228-230.
Doonan, T. J. 1986. A demographic study of an isolated population of the gopher
tortoise, Gopherus polyphemus; and an assessment of a relocation procedure for
tortoises. M. S. Thesis. University of Central Florida, Orlando.
Douglass, J. F., and C. E. Winegamer. 1977. Predators of eggs and young of the gopher
tortoise Gopherus polyphemus (Reptilia, Testudines, Testudinidae) in southern
Florida. Journal of Herpetology 11:236-238.


39
submucosal lymphoid hyperplasia were seen in the glottis of one tortoise. The hyperplasia
extended into the cranial tracheal epithelium. In that tortoise, there also were multifocal
areas of epithelial cell hyperplasia in the lung. Five other diseased tortoises had focal to
multifocal lymphoid aggregates in the lung interstitia.
The gastrointestinal tracts tended to have increased numbers and larger lymphoid
aggregates in the submucosa compared to those of clinically healthy tortoises. In one
tortoise there was mucous cell hyperplasia of the colon and in another there was a severe
colitis with basal epithelial cell hyperplasia and submucosal lymphoid hyperplasia, with
infiltrat es of large numbers of heterophils. Four other tortoises had increased lymphoid
aggregates in the esophagus, stomach and/or small intestine. Three additional tortoises
had increased lymphoid aggregates in the submucosa of the cloaca.
The kidneys of all diseased tortoises contained golden brown pigment granules
within renal epithelial cells. Hepatocytes of most tortoises contained similar granules, and
pathologic changes were seen in the livers of nine tortoises. In seven, there were
increased numbers and size of melanomacrophages in the liver and increased amounts of
golden brown granules. One tortoise had cuffing of the central vein by lymphocytes and
heterophils; another had aggregates of lymphocytes and melanomacrophages.
Electron micrographs of tissues of one tortoise from Sanibel Island and the tortoise
from Indian River County demonstrated organisms consistent with Mycoplasma on the
surface of the nasal mucosa (Figure 3-5). Associated epithelial cells had vacuolated
cytoplasm and inflammatory cell infiltrates were present in the mucosa. Increased
numbers of mucous epithelial cells were seen, consistent with light microscopic findings.


61
Figure 5-1. Flow chart showing initial distribution of seropositive, clinically ill, culture
and PCR positive tortoises.


69
urinated), or having one suspect culture or PCR result recorded as having been infected by
horizontal transmission. The four classes were 1) ELISA, clinically, and culture/PCR
negative (-/-/-), 2) ELISA positive, clinically and culture/PCR negative (+/-/-), 3) ELISA
and clinically positive, and culture/PCR negative (+/+/-), and 4) ELISA, clinically, and
culture/PCR positive (+/+/+). The first data set (Table 5-2) consisted of the following
observations (class, yes, no): 1,0,5; 2,0,2; 3,2,2; and 4,4,0. The second set (Table 5-3),
for the last 14 mo, was comprised of the following observations: 1,0,4; 2,0,1; 3,0,1; and
4,5,0. The last, cumulative, data set, included the following observations: 1,0,5; 2,0,3;
3,2,3; and 4,9,0. Chi-square tests of association of exposure status and transmission were
significant for all data sets, with 3 degrees of freedom (df) and P < 0.001 for all sets
(Tables 5-2, 5-3, 5-4).
Table 5-2. Contingency table for exposure status and transmission occurrence for the first
10 mo (August 1994 June 1995) of the gopher tortoise URTD pairing study.
Exposure Status
ELISA/Clinical/Culture or PCR
Transmission
Yes No
-/-/-
0 5
+1-1-
0 2
+/+/-
2 2
+/+/+
4 0
X2, 3 df= 10.8, P = 0.013, power = 0.80
Table 5-3. Contingency table for exposure status and transmission occurrence for the last
14 mo (June 1995 July 1996) of the gopher tortoise URTD pairing study.
Exposure Status
ELISA/Clinical/Culture or PCR
Transmission
Yes No
-/-/-
0 4
+1-1-
0 1
+1+1-
0 1
+1+1+
5 0
X2, 3 df = 11.0, P = 0.012, power = 0.81


91
A clinically healthy tortoise (i.e., one with no nasal or ocular discharge or other
signs) may have positive culture and PCR results, with either positive or negative ELISA
results. It may be in the early stages of infection or recrudescence. In the former case, the
ELISA value may be negative, but should rise within 6 wk. In the latter case, the ELISA
value may be quite high. Such tortoises may be capable of transmitting the mycoplasma
under the appropriate conditions. Although direct contact (nose-to-nose) seems to be the
most important route, transmission through water or on food cannot be ruled out. When
tortoises drink, they often expel water through their noses for short distances, up to 50
cm, or sneeze forcefully after drinking, aerosolizing the contents of the nasal passages. If
tortoises are in close contact with one another, spatially or temporally, such occurrences
may allow transmission of M. agassizii. Tortoises with slight nasal discharge, virtually
undetectable, can also aerosolize bacteria by sneezing.
Although long-term studies on the effect of URTD on survival of individuals and
populations have not been conducted, the evidence from surveys of desert tortoise
populations and from Sanibel Island indicate that the disease can have severe negative
impacts on population viability. Declines of 25-50% over 1-3 yr, and of 30-90% over 10
yr, are catastrophic for species that take 10-20 yr to reach maturity and have recruitment
rates of 1-2%. Without marked improvement in recruitment rates, affected populations
are unlikely to recover within a reasonable time frame.
Developing Questions and Conducting Surveys or Monitoring Programs
Third, questions related to management goals, and taking into account test
interpretation and tortoise behavior, must be developed. Some of the most common


13
Goats can suffer 60-100% mortality when infected by M. mycoides subsp.
mycoides, M. mycoides capri, or Mycoplasma strain F38, making contagious caprine
pleuropneumonia the most economically important goat disease. Pathologically, signs and
lesions are similar to those seen in CBPP, with the addition of polyarthritis (Cottew 1984).
Humans are susceptible to infection with M. pneumoniae, which causes
tracheobronchitis and, less commonly, primary atypical, or walking, pneumonia (Chanock
et al. 1963, Clyde 1983). The upper respiratory tract is affected, not the alveoli, and otitis
media is seen also (Clyde 1979, Mansel et al. 1989). Immunopathological sequelae,
although rare, can include arthritis, dermal lesions, cardiopathy, and neurological
complications (Johnston et al. 1983, Lyell et al. 1967, Naftalin et al. 1974, Murray et al.
1975). Histologically, perivascular and peribronchial monocytic infiltrations with some
neutrophilic exudate are seen (Dajani et al. 1965). Most commonly affected are 5-9 yr old
children, with a decline in incidence until about age 25, then an increase in 30-40 yr olds
(Denny et al. 1971). The disease is endemic, with slight increases in late summer and fall,
and cyclic epidemics occurring at 4-7 yr intervals (Krause and Taylor-Robinson 1992).
Mycoplasmal Diseases of Wildlife
Research on the impact of mycoplasmosis on wildlife is limited, but recent
developments are provoking interest. In early 1994, house finches (Carpodacus
mexicanus) in the mid-Atlantic and northeastern regions of the United States were found
with conjunctivitis, rhinitis, and sinusitis. Many birds were found dead or dying and were
submitted to wildlife care facilities, veterinary hospitals, and state agricultural laboratories.
Many of those birds had lesions that were histologically compatible with mycoplasma


Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the Requirements
for the Degree of Doctor of Philosophy
UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES, GOPHERUS
POLYPHEMUS: PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION, AND
IMPLICATIONS FOR CONSERVATION AND MANAGEMENT
By
GRACE SHERYL MCLAUGHLIN
May 1997
Chair: Donald J. Forrester, Ph. D.
Cochair: Mary B. Brown, Ph. D.
Major Department: Wildlife Ecology and Conservation
Upper respiratory tract disease (URTD) of tortoises is caused by the molhcute
Mycoplasma agassizii, and is characterised by nasal and ocular discharge, palpebral
edema, and conjunctivitis. Hyperplasia and dysplasia of the nasal passage and cavity
epithelia and inflammatory infiltrates are seen histologically. In order to provide data for
management decisions and to better understand URTD, I studied uninfected, naturally
infected, and experimentally infected tortoises. The pathological and immune responses of
tortoises to and transmission of M. agassizii were investigated using clinical and
histological observations, culture and polymerase chain reaction (PCR) tests, and an
enzyme-linked immunosorbent assay (ELISA). Infection with M. agassizii caused mild to
severe damage to mucosal and olfactory nasal epithelia, with increased damage in longer-


I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and quality,
as a dissertation for the degree of Doctor of Philosophy.
Donald J. Forrester, Chair
Professor of Wildlife Ecology and Conservation
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Yny ^ &OOW
Mary B. Brown, Cochair
Associate Professor of Veterinary Medicine
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Melvin E. Sunquist
Associate Professor of Wildlife Ecology
and Conservation
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Wiley M. Kitchens
Associate Professor of Wildlife Ecology
and Conservation
I certify that I have read this study and that in my opinion it conforms to acceptable
standards of scholarly presentation and is fully adequate, in scope and quality, as a
dissertation for the degree of Doctor of Philosophy.
Elliott R. Jacobson
Professor of Veterinary Medicine


103
Kerr, K. M, and N. O. Olson. 1967. Pathology in chickens experimentally inoculated or
contact-infected with Mycoplasma gcillisepticum. Avian Diseases 11: 559-578.
Kirchhoff, H., R. Schmidt, H. Lehmann, H. W. Clark, and A. C. Hill. 1996. Mycoplasma
elephantis sp. nov., a new species from elephants. International Journal of Systematic
Bacteriology 46: 437-441.
Kleven, S. H., H. H. Fan, and K. Turner. 1996. Displacement of virulent Mycoplasma
gallisepticum in chickens with Uve vaccines. IOM Letters 4: 284-285.
Knowles, C. 1989. A survey for diseased desert tortoises in and near the Desert Tortoise
Natural Area, Spring 1989. Report prepared for the Bureau of Land Management,
Riverside, California, Contract No. CA 950-(T9-23), 26 pp.
Krause, D. C., and D. Taylor-Robinson. 1992. Mycoplasmas which infect humans. Pp.
417-444 in: Maniloff, I, R. N. McElhaney, L. R. Finch, J. B. Baseman (eds.).
Mycoplasmas: molecular biology and pathogenesis. American Society for
Microbiology, Washington, D. C.
Kume, K_, Y. Kawakubo, C. Morita, E. Hayatsu, and M. Yoshioka. 1977.
Experimentally induced synovitis of chickens with Mycoplasma synoviae: effects of
bursectomy and thymectomy on course of infection for the first four weeks. American
Journal of Veterinary Research 38: 1595-1600.
Lai, W. C., M. Bennett, and S. P. Pakes. 1996. Successful immunization of mice after
infection with virulent Mycoplasma pulmonis. IOM Letters 4: 52.
Landers, J. L., and D. W. Speake. 1980. Management needs of sandhill reptiles in
southern Georgia. Proceedings of the Annual Conference Southeastern Association
of Fish and Wildlife Agencies 34: 515-529.
Landers, J. L., J. A. Gamer, and W. A. McRae. 1980. Reproduction of the gopher
tortoise (Gopherus polyphemus) in southwestern Georgia. Herpetologica
36:353-361.
Landers, J. L., W. A. McRae, and J. A. Gamer. 1982. Growth and maturity of the
gopher tortoise in southwestern Georgia. Bulletin of the Florida State Museum,
Biological Sciences 27:81-110.
Lawrence, K., and J. R. Needham. 1985. Rhinitis in long-term captive Mediterranean
tortoises (Testudo graeca and T. hermanni). Veterinary Record 117: 662-664.
Levell, J. P. 1995. A field guide to reptiles and the law. Serpents Tale, Excelsior, MN.
240 pp.


9
(ELISA) to detect antibodies against the mycoplasma in plasma and serum samples was
developed (Schumacher et al. 1993), and experiments were undertaken to fulfill Kochs
postulates. The disease was induced by inoculation of tortoises with pure cultures of the
mycoplasma, but not Pasteurella testudinis (Brown et al. 1994). Histologically, the
lesions were consistent with those seen in the previously examined naturally infected
tortoises. Additional work led to the development of a polymerase chain reaction (PCR)
test to detect the bacteria in nasal flush and swab samples (Brown et al. 1995).
Histologic examination of nasal tissues and microbiologic evaluation of flush and
swab samples from clinically healthy and clinically ill tortoises from Las Vegas Valley,
Nevada, resulted in findings in the ill tortoises similar to those in the California tortoises,
with 92% showing lesions of URTD, 50% being culture positive for M. agassizii, and
100% reacting positively in the ELISA (Jacobson et al. 1995). However, 73% of the
healthy tortoises had lesions consistent with URTD, 50% were culture positive for M
agassizii, and 42% were seropositive for antibodies against the bacteria. These findings
demonstrate that the disease can exist in a subclinical form in a substantial proportion of a
population. An annual cycle of convalescence and recrudescence of clinical signs has been
seen in captive desert (I. M. Schumacher, pers. comm.) and gopher (D. L. Morris, pers.
comm.) tortoises. Other mycoplasmal diseases also can exist as chronic, subclinical
infections, with recurrence of clinical signs and increases in transmission potential when
the host is stressed (see below). Annual fluctuations in temperature, rainfall, and forage
availability may be sufficient to cause detectable outbreaks in an infected population.
Increased morbidity and mortality may occur in times of unusually severe environmental
stress, such as prolonged drought, hurricanes, excessive rainfall with flooding of burrows,


3
and forbes, and readily ingest the fruits of many shrubs when available (Gamer and
Landers 1981, Macdonald and Mushinsky 1988). Subsequent tortoise movements may
help spread the seeds to suitable habitat, and some seeds (e.g., gopher apple) may
germinate more readily after passage through the tortoise gut.
Gopher tortoises are long-lived, slow to mature, and have a low reproductive rate.
Age estimates extend up to 100 yr, although 60-80 yr is considered a more reasonable
estimate. Age to sexual maturity ranges from 10 to 20 yr, with possible latitudinal
association (Alford 1980, Iverson 1980, Landers et al. 1982, Wright 1982, Doonan 1986,
McLaughlin 1990, Mushinsky et al. 1994). Gopher tortoises lay one clutch of eggs
annually, generally ranging from 2-10 eggs in size, with the average being 4-8 (Dietlein
and Franz 1979, Iverson 1980, Landers et al. 1980, Linley and Mushinsky 1994). In some
areas with excellent food resources and large tortoises, clutch sizes may average 9-10,
with a range of 6-14 eggs (McLaughlin, 1990). Although some wild individuals have
produced over 20 eggs in one clutch (Godley 1989, L. Macdonald, personal
communication), this is very unusual. With predation rates of up to 95% on eggs and
hatchlings, and further high losses of juveniles aged 1-5 yr, less than 10%, and possibly as
low as 1%, of eggs laid eventually produce reproductive adults (Douglass and Winegamer
1977, Auffenberg and Iverson 1979, Alford 1980, Iverson 1980, Landers et al. 1982,
Diemer 1986, Wilson 1991).
The gopher tortoise is listed in Appendix II of the Convention on International
Trade in Endangered Species of Wild Fauna and Flora (C.I.T.E.S.), which requires
permits for the exportation of the species from the U. S. to any signatory nation, or for re
exportation (Levell 1995). However, if federal or state regulations are more restrictive


64
Results
General Observations
Observations indicated varying degrees of interaction between pair members.
Several males were excluded from the original burrow in the pen by their female partners.
Until one of the two dug a new burrow, those males, particularly 125, 185, 201, and 235,
spent most of the day in a comer of the pen, under the shade cloth, and sometimes spent
the night out of the burrow. Other pairs spent considerable time in courtship, or foraging
in close proximity to one another.
In March, 1995, five hatchling tortoises were found in pen Dl. The female (135)
may have been gravid or storing sperm when she arrived at the UF facility in April 1994,
and laid the clutch during the summer of 1994. Alternatively, she may have copulated
with the male (241) and ovulated shortly after the pairing in August 1994. Based on
gopher tortoise physiology (Taylor 1982), however, I believe the former explanation is
more likely. One hatchling was removed and euthanatised due to congenital
abnormalities; the other four were left in the pen.
Three tortoises were euthanatised in June 1995 (Table 5-1). One, number 287
(pen DIO), was an initially seropositive, clinically ill, culture and PCR negative female
whose partner (125) had not become ill. She failed to reproduce, and a large mass could
be seen on radiographs. At necropsy, a large urolith (bladder stone), approximately 5 cm
in diameter, was removed. The second, number 213 (pen D5), was an initially
seropositive, clinically ill, culture and PCR positive female whose partner had become ill
within one month of pairing. She failed to lay six eggs, and on exploratory surgery it was
discovered that both oviducts had ruptured and the eggs were free in the coelomic cavity.


82
Discussion
Unlike various mycoplasmal infections of rodents and poultry (Simecka et al.
1992) and wild ducks (Goldberg et al. 1995), there was no evidence to support a
hypothesis of vertical transmission of M. agassizii. Therefore, it should be possible to
collect eggs from infected female tortoises, incubate the eggs, and release the hatchlings
with no risk of spreading the disease.
Vitellogenesis begins in August or September and continues to December (Taylor
1982), providing an extensive period for deposition of antibodies. In spite of that, the
antibody level in egg yolks and hatchling plasma was only 10 -20% of that in the maternal
plasma. Thus, there is no evidence that female gopher tortoises are sequestering
antibodies in the egg components. This is in contrast to birds, particularly chickens, which
deposit large amounts of antibodies in their eggs in a very short time period. Antibody
levels in chicken eggs can be several orders of magnitude greater than that in the maternal
plasma.
Hatchling antibody levels decline slowly over the first year of life, as the maternal
antibodies are broken down (I. M. Schumacher, unpub. data). This process is much
slower in tortoises than in mammals and birds, where such passively acquired antibodies
decline within weeks or a few months after birth or hatching. It is not known if antibodies
against M agassizii affect juvenile responses to infection with the organism Although
adult tortoises that have developed an antibody response against M. agassizii respond very
quickly and adversely to subsequent exposure (Chapter 4), that response may be mediated
more by cellular immune components (e g., macrophages and heterophils) than by


49
Results
Clinical Signs
Of the six control group tortoises, none showed consistent clinical signs nor had
positive culture, PCR, or ELISA results. At each of three time points, one control
tortoise exhibited mild ocular signs that were probably associated with environmental
conditions. One of two sentinels developed clinical signs at 12 wk PI, although she had
not been inoculated with culture or medium. Of the naive tortoises, 67% exhibited clinical
signs beginning 2-3 wk PI, 79% were clinically ill by 8 wk PI, and 94% had shown signs
by 16 wk PI (Table 4-1). One naive tortoise never exhibited clinical signs. All seven
challenged tortoises developed clinical signs of URTD before dormancy. Five (71%)
challenged tortoises and one naive tortoise exhibited clinical signs soon after emergence.
No tortoises became moribund or died during the study.
At 2 wk PI, six challenge and 17 naive tortoises were examined. Challenged
tortoises had higher total clinical sign scores than naive tortoises (2.6 vs. 0.5, P < 0.001),
and higher scores for nasal discharge (1.5 vs. 0.1, P < 0.001), ocular discharge (0.8 vs. 0,
P<0.02), and palpebral edema (1.5 vs. 0.6, P < 0.02) (Table 4-1). Significantly more
challenge than naive tortoises exhibited nasal and ocular discharge (67 vs. 6%, P < 0.01;
50 vs. 0%, P < 0.02; respectively) (Figure 4-1). No consistent differences were seen
between the two groups at later sampling times (Figure 4-2).


60
Experimental Design
Fifteen pairs consisting of one male and one female were established in August,
1994. Due to the limited availability of seropositive and/or clinically ill tortoises, the
sample design was not balanced. Five pairs of asymptomatic, ELISA-, PCR-, and culture
negative control tortoises were established as controls. The other ten pairs consisted of a
clinically healthy, ELISA-, PCR-, and culture-negative tortoise that had been at UF since
April, 1994 (resident), and an ELISA-positive or -suspect (irrespective of PCR or culture
status) tortoise of the opposite sex and similar size (Figure 5-1, Table 5-1). One resident
female was paired with an ELISA-negative, but clinically ill, culture- and PCR-positive
male. The serosuspect and two seropositive tortoises were clinically healthy, and had
negative culture and PCR results. The remaining six seropositive tortoises showed
moderate clinical signs of illness. Three of those six were culture- and PCR-negative, one
was culture- and PCR-positive, one was culture-positive but PCR-negative, and one was
culture-negative and PCR-positive.
Behavioral observations and clinical signs of URTD were recorded
opportunistically (see Chapter 2), and blood and nasal flush samples were collected from
all tortoises in August and October, 1994, and in March and August, 1995. In 1996,
samples were collected in February or March, and in July or August. Clinical signs and
weights were recorded, and photographs were taken at each sampling time. Data were
analysed using Chi-square and Fisher Exact tests for differences in proportions of tortoises
becoming infected under different exposure conditions. Infected animals were
euthanatized in July, 1996, except as noted below, and uninfected tortoises were released
pursuant to Florida Game and Fresh Water Fish Commission permit number WR96224.
(


70
Table 5-4. Contingency table for exposure status and transmission occurrence for the
duration (August 1994 June 1995) of the gopher tortoise URTD pairing study.
Exposure Status
ELISA/Clinical/Culture or PCR
Transmission
Yes No
-/-/-
0 5
+/-/-
0 3
+/+/-
2 3
+/+/+
9 0
X2, 3 df = 17.2, P = 0.0006, power = 0.96
Table 5-5. Transmission probabilities and tests of significant differences of proportions
for different exposure status classes of tortoises relative to upper respiratory tract disease.
Exposure
Transmission
Probability of
Fisher Exact
Status
Yes
No
Transmission
Test, P value
ELISA
negative
0
5
0
positive
11
6
0.65
0.018
Clinical illness
negative
0
8
0
positive
11
3
0.79
0.001
Culture / PCR
negative
2
11
0.15
positive
9
0
1.00
0.0002
Because transmission was defined very conservatively for the purposes of this
study, two tortoises were categorized as having been infected via horizontal transmission
based solely on the appearance of mild clinical signs. Neither animal seroconverted nor
had positive or suspect culture or PCR results during the second half of the study. Both
tortoises were housed with clinically ill, seropositive tortoises that were culture and PCR
negative. If those tortoises were designated negative relative to transmission, the final


LITERATURE CITED
Alford, R. A. 1980. Population structure of Gopherus polyphemus in northern Florida.
Journal of Herpetology 14: 177-182.
Anonymous. 1989. National poultry improvement plan and auxiliary provisions. U. S.
Department of Agriculture, Animal and Plant Health Inspection Service 91-40,
Hyattsville, MD. 89 pp.
Auffenberg, W., and R. Franz. 1982. The status and distribution of the gopher tortoise
(Gopherus polyphemus). Pp. 95-126 in R. B. Bury (ed.). North American tortoises:
conservation and ecology. U. S. Fish and Wildlife Service, Wildlife Research Report
12.
Auffenberg, W., and J. B. Iverson. 1979. Demography of terrestrial turtles. Pp. 541-569
in M. Harless and H. Morlock (eds.). Turtles: perspectives and research. John Wiley
and Sons, New York.
Baskerville, A. 1972. Development of the early lesions in experimental enzootic
pneumonia of pigs: an ultrastructural and histological study. Research Veterinary
Science 13: 570-578.
Behymer D., D. Jessup, K. Jones, C. E. Franti, H. Riemann, and A. Bahr. 1989.
Antibodies to nine infectious disease agents in deer from California. Journal of Zoo
and Wildlife Medicine 20: 297-306.
Berry, K. H. 1990. The status of the desert tortoise in California in 1989. Annual report.
Bureau of Land Management, Riverside, California. 94 pp.
Berry, K. H. in press. Demographic consequences of disease in two desert tortoise
populations in California, USA. Proceedings: conservation, restoration, and
management of tortoises and turtlesan international conference. WCS Turtle
Recovery Program and the New York Turtle and Tortoise Society.
Black S. R., I. K. Barker, K. G. Mehren, G. J. Crawshaw, S. Rosendal, L. Ruhnke, J.
Thorsen, P. S. Carman. 1988. An epizootic o Mycoplasma ovipneumoniae infection
in captive Dali's sheep (Ovis dalli dalli). Journal of Wildlife Diseases 24: 627-635.
97


22
A 2-3 ml blood sample was drawn from the jugular or brachial vein and placed in a
lithium heparin Vacutainer tube (Becton Dickinson and Company, Rutherford, New
Jersey). Blood was centrifuged and an aliquot of plasma was removed for antibody
screening by an enzyme-linked immunosorbent assay (ELISA) (see below).
After cleansing the area around the nares with alcohol dampened gauze, nasal flush
samples were collected by flushing with approximately 0.5 ml sterile SP4 broth using a 1
ml syringe without needle. Calcium alginate-tipped swabs were inserted gently into the
nares, and a sample was obtained and streaked onto SP4 agar plates.
Culture Procedures
A 100 pi aliquot of the flush sample was used for polymerase chain reaction (PCR)
analysis; the remaining sample was serially diluted ten-fold to 102 and incubated at 30C
for a maximum of three weeks, or until determined to be positive or contaminated.
Twenty pi of each dilution were placed on SP4 agar and incubated at 30C and 5% CO, as
were the streak plates. Plates were examined regularly for a maximum of six weeks to
detect the growth of mycoplasma. In the second year of the study, broth cultures were
incubated for 24-48 hr before removing the aliquot for PCR, as that modification increased
the sensitivity of the PCR (G. S. McLaughlin and D. R. Brown, unpub. data).
PCR Procedure
Nasal aspirate samples were analyzed for the presence of Mycoplasma agassizii
DNA based upon PCR amplification of the 16S rRNA gene (Brown et al. 1995). Nasal
flush and culture samples were centrifuged for 60 min at 14,000 rpm at 4C, and the


47
Methods
Acquisition of tortoises, intake and husbandry procedures, clinical assessments,
sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed
in Chapter 2.
Statistical Analyses
The onset and severity of clinical signs of URTD, and ELISA, PCR, and culture
results for the challenge tortoises were compared to those for control and naive tortoises
using the SAS system (SAS Institute, 1988). Data from an additional naive animal
infection experiment were included after it was determined that the data did not differ
significantly from those collected from naive animals in this study. This ensured large
enough sample sizes for meaningful comparisons at more time points. Differences in the
severity of clinical signs, histologic lesions, and ELISA data among the three treatment
groups were compared by an analyses of variance-type logistic regression using maximum
likelihood estimators to compensate statistically for the different number of tortoises in
each group at each sampling date. Percentages of tortoises showing clinical signs at
different time points were compared by Fishers Exact test, with a P value of 0.05
accepted as significant.
Experimental Design
Four groups of tortoises were established. Three groups [control {n = 6), naive (n
= 11), and sentinel (n = 2)] had no history of exposure to M. agassizii, while the fourth, or
challenge group (n = 8), had previous history of exposure as indicated by a positive result


66
Figure 5-2. Burrow map showing interconnections among pens Dl, D6, D7, and D8;
between D3 and D4, and between D9 and DIO. Burrow entrances.
Evidence of Transmission of Mycoplasma agassizii
The male control tortoise (147) became ill, seroconverted, and had positive culture
and PCR results (Table 5-1). His partner became cbnically ill and seroconverted after he
returned to his original pen. No other control tortoises became ill or seroconverted. The
D-section burrowing activities mentioned above resulted in the exposure of all tortoises in
pens Dl and D7, which until that time had been clinically healthy, to clinically ill tortoises
with positive culture and/or PCR results. Three became clinically ill, and had increased
ELISA readings, with one of two initially negative tortoises seroconverting. Those
activities also resulted in the exposure of the tortoises in pen D8 to cbnically ill, culture
and PCR positive tortoises. The seronegative male (130) became ill, had positive culture
results, and seroconverted; the cbnically ill, seropositive female (257), which had been
culture and PCR negative until that time, also became culture positive. All four hatchlings
in pen Dl became severely ill, seroconverted, and had positive culture results. All were
euthanatised, and pure cultures of M. agassizii were recovered from the nasal cavities and
conjunctiva on necropsy.


90
80
70
60
50
40
30
20
10
0
4-
iNaive
CS Total ND OD
ED
CJ
Percent of gopher tortoises infected with Mycoplasma agassizii positive for any clinical sign and each individual sign at 2
weeks postinfection. Tortoises in the challenge group had serological evidence of prior exposure to M. agassizzi, but
those in the naive group did not. CS = clinical signs, ND = nasal discharge, OD = ocular discharge, ED = palpebral
edema, CJ = conjunctivitis. Indicates significant difference between groups, P < 0.05.



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UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES, GOPHERUS POLYPHEMUS: PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION, AND IMPLICATIONS FOR CONSERVATION AND MANAGEMENT BY GRACE SHERYL MCLAUGHLIN A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNTVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 1997

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DEDICATION This dissertation, the years that went into it, and the drive for knowledge that underpinned it, are a result of, and dedicated to, my parents: Grace Slater Gibson "Billie" McLaughlin 1928-1978 and Robert Douglas McLaughlin 1925-1994 Thank you.

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ACKNOWLEDGMENTS I thank Dr. Don Forrester for accepting me as a graduate student, and for his support and guidance through the years. Drs. Kathy Ewel and Paul Gibbs assisted my growth scientifically and professionally in the first half of my program Dr. Ewel's support was instrumental in obtaining my fellowship, and Dr. Gibbs was responsible for a trip to Australia. Drs. Mel Sunquist and Wiley Kitchens weathered the changes in my project with grace and humor, and Dr. Kitchens was especially helpful in teaching me to argue my positions and not back down when I knew I was right. It took several years for Dr. Mary Brown to get me into her lab, and her support in presenting me to her colleagues is appreciated. Dr. Elliott Jacobson has done his best to teach me clinical pathology and histopathology and has been very supportive of my contributions to the overall project. Dr. Paul Klein has given me some valuable insights into the critical thinking process. Without my co-workers Drs. Dan Brown and Isabella Schumacher, Sylvia Tucker, Barbara Crenshaw, and Cathie McKenna, and technicians Alyssa Whitemarsh, Michael Lao, and Dave Bunger, this research would have been impossible. I benefited from Dan's, Isa's and Barb's teaching abilities, and their willingness to discuss theory, practical applications, philosophical underpinnings and differences of opinion. Mr. Clement Lindsey and his staff cared for my research animals. I thank Dr. Tim Gross for all his help with eggs and hatchlings, and John Wiebe and Carla Weiser for their care. Drs. Dale Jackson and Michael Ewert provided advice. Drs. Bruce Homer and Claus Buergelt have given me more strength and support than they iii

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knew and taught me much. Many of the VMTH personnel, especially Neil, Danielle, Tom, Dana, and An have assisted and taught me, and cheered me up when I needed it. Ms. Joan Berish has provided an incredible amount of advice, support, and friendship throughout the eight years I have been studying gopher tortoises, and I only hope to reciprocate. Garry Foster's assistance through my early years at UF were invaluable, as were the support of my fellow graduate students in Dr. Forrester's lab, Marisol Sepuh/eda and Don Coyner. My brother Mark, my "sister" Megan, and friends Vicky, Bill, Sharon, Kay, Pierre, Andrea, and Tania helped me get here and stick it out, and deserve thanks for their support. There have been several other people who have helped me at various times and in different ways, especially those of the gopher tortoise sodality, and my thanks go to them also. I also thank the members and friends of the Unitarian Unfversalist Fellowship of Gainesville for being my family and caring for me. I was supported by a Presidential Research Fellowship and a Gatorade Grant from the University of Florida, and funds from The Walt Disney World Company. iv

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TABLE OF CONTENTS page ACKNOWLEDGMENTS iii ABSTRACT viii CHAPTERS 1 INTRODUCTION 1 Gopher Tortoise Natural History 1 Tortoises and Upper Respiratory Tract Disease 6 Mycoplasmal Respiratory Diseases in Domestic Animals and Humans 10 Mycoplasmal Diseases of Wildlife 13 Chronic Manifestations of Mycoplasmal Infections 18 Project Overview and Specific Objectives 18 2 METHODS 21 Tortoises, Intake Procedures, Clinical Assessments and Sampling Methods 21 Culture Procedures 22 PCR Procedure 22 ELISA Procedure 24 Study Group Assignment 25 Husbandry Procedures 26 Necropsy Procedures 27 Histopathology Procedures 28 Statistical Analyses 29 3 NATURALLY OCCURRING UPPER RESPIRATORY TRACT DISEASE 30 Methods 30 Tortoises 30 Necropsy and Histology Procedures 31 Microbial Isolation 31 Electron Microscopy 32 Results 33 Normal Anatomy and Histology 33 Pathologic Findings 37 ELISA and PCR Results 40 Microbial Isolation Results 40 Discussion 40 v

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4 EFFECTS OF REPEATED EXPOSURE ON SEROPOSITIVE ADULTS 46 Introduction 46 Methods 47 Statistical Analyses 47 Experimental Design 47 Results 49 Clinical Signs 49 Culture and PCR Results 53 ELISA Results 53 Histology Results 55 Discussion 55 5 HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII . . 59 Introduction 59 Methods 59 Experimental Design 60 Results 64 General Observations 64 Evidence of Transmission of Mycoplasma agassizii 66 Transmission Probabilities 68 Discussion 71 6 VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII 75 Introduction 75 Methods 75 Egg Collection and Incubation 76 Culture and PCR Procedures 76 ELISA Procedures 77 Results 78 Clutch Sizes, Fertility and Hatching Rates 78 Culture and PCR Results 79 ELISA Results 80 Discussion 82 7 ENVIRONMENTAL TRANSMISSION OF MYCOPLASMA AGASSIZII 84 Introduction 84 Methods 84 Results 85 Discussion 86 vi

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8 CONSERVATION AND MANAGEMENT IMPLICATIONS OF UPPER RESPIRATORY TRACT DISEASE 88 Implications for Conservation and Management 88 Establishing Goals 89 Understanding URTD and Test Results 89 Developing Questions and Conducting Surveys or Monitoring Programs 91 Weighing Management Options and Formulating Management Plans 93 Summary of Conservation and Management Implications 95 Further Research 96 LITERATURE CITED 97 BIOGRAPHICAL SKETCH 110 vii

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES, GOPHERUS POLYPHEMUS: PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION, AND IMPLICATIONS FOR CONSERVATION AND MANAGEMENT By GRACE SHERYL MCLAUGHLIN May 1997 Chair: Donald J. Forrester, Ph. D. Cochair: Mary B. Brown, Ph. D. Major Department: Wildlife Ecology and Conservation Upper respiratory tract disease (URTD) of tortoises is caused by the molhcute Mycoplasma agassizii, and is characterised by nasal and ocular discharge, palpebral edema, and conjunctivitis. Hyperplasia and dysplasia of the nasal passage and cavity epithelia and inflammatory infiltrates are seen histologically. In order to provide data for management decisions and to better understand URTD, I studied uninfected, naturally infected, and experimentally infected tortoises. The pathological and immune responses of tortoises to and transmission of M. agassizii were investigated using clinical and histological observations, culture and polymerase chain reaction (PCR) tests, and an enzyme-linked immunosorbent assay (ELISA). Infection with M. agassizii caused mild to severe damage to mucosal and olfactory nasal epithelia, with increased damage in longerviii

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term infections. Although clinically healthy, ELISA-positive, culture and PCR-negative tortoises may have eliminated the bacteria, when five such animals were examined at necropsy, three were found to harbor M. agassizii in the nasal cavities. When seropositive tortoises were challenged with M. agassizii, a more rapid and more severe clinical response resulted than on initial exposure, and plasma antibody levels began rising more quickly. When uninfected animals were housed with infected individuals, horizontal transmission occurred, probably via direct contact, but possibly via food, water, or fomites. Transmission was more likely to occur from a tortoise that was clinically ill and culture or PCR-positive. There was no discernible transmission when tortoises inhabited pens or entered burrows previously occupied by ill tortoises. There was no demonstrable vertical transmission, although there was transfer of maternal antibodies via egg yolk. The level of antibodies in egg yolk or hatchling plasma was approximately 10-20% of that in maternal plasma. Movement of tortoises during relocation, repatriation, or restocking efforts potentially could transport M. agassizii to previously uninfected sites. Because of the uncertainty involved in determining latent infections, ELISA-positive animals should not be moved to locations with no seropositive individuals. However, they can be used in captive breeding efforts and their offspring released into the wild. Clinically ill animals should not be relocated to new sites, and should not be housed with clinically healthy animals in temporary holding situations, such as on-site relocations, or in captive breeding programs. ix

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CHAPTER 1 INTRODUCTION Gopher Tortoise Natural History Gopher tortoises, Gophems polyphemns, are found in the southeastern United States, on the coastal plain from southern South Carolina south through Georgia and throughout Florida, and west through southern Alabama, Mississippi, and Louisiana. The major population concentrations are in Florida and southern Alabama and Georgia, with only remnant populations in South Carolina, Mississippi and Louisiana (Auffenberg and Franz 1982, Diemer 1992a). Populations are concentrated in areas with deep sandy soils suitable for digging. Vegetation associations in which tortoises are found include longleaf pine-xerophytic oak woodlands, palmetto scrub, sand pine scrub, oak scrub, beach scrub, coastal strands, pine flatwoods, dry prairies, native pasture, and savanna, as well as ruderal habitats (Landers and Speake 1980, Lohoefener and Lohmeier 1981, McRae et al. 1981, Campbell and Christman 1982, Diemer 1986, Breininger et al. 1988). 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 (Eisenberg 1983). Gopher tortoises live in loose colonies, with considerable movement of tortoises among groups over the years (Diemer 1992b). Colonies may be defined more by the availability of suitable soils for digging burrows, or the distribution of food resources, than by social

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interactions (Campbell and Christman 1982). Gophers are the most fossorial of the four North American species of tortoises, digging burrows that may extend 5 meters down from the surface and 15 meters in length (Hansen 1963, Diemer 1986). The burrows provide a microclimatically stable environment for not only the tortoises, but also for numerous commensals. Approximately 60 vertebrate species—from snakes to birds—and over 300 invertebrates— including spiders, crickets, and beetles— have been found in tortoise burrows or observed using them as permanent homes or refuges from heat, cold, fire, and predators (Jackson and Milstrey 1989, Lips 1991, Witz et al. 1991). Some invertebrate species are obligate commensals, and occur only in active tortoise burrows, where they feed on tortoise feces and other invertebrates and, in turn, are eaten by other commensals, such as gopher frogs (Rana areolata) and Florida mice (Podomys floridanus) (Woodruff 1982). Several species that exclusively or frequently use tortoise burrows have legal protection in Florida and other parts of their ranges. These include scarab beetles (F. Scarabaeidae), indigo {Drymarchon corais couperi) and pine (Pituophis melanoleucus) snakes, gopher frogs, mole skinks (Eumeces egregius), burrowing owls (Athene cunicularia floridana), and Florida mice (Cox et al. 1987). The soil disturbance resulting from the digging of the burrows allows deeper access of air and water into the soil profile, as well as providing bare mineral soil patches on the surface. When a burrow is abandoned the soil mound, or apron, in front of the entrance no longer undergoes continual disturbance, allowing certain plants to colonize the area. The composition of the plant assemblage on the mound may differ from that in the surrounding undisturbed area, providing a mosaic of small patches in the habitat (Breininger et al. 1988, McLaughlin 1990). Tortoises consume a wide variety of grasses

PAGE 12

and forbes, and readily ingest the fruits of many shrubs when available (Garner and Landers 1981, Macdonald and Mushinsky 1988). Subsequent tortoise movements may help spread the seeds to suitable habitat, and some seeds (e.g., gopher apple) may germinate more readily after passage through the tortoise gut. Gopher tortoises are long-lived, slow to mature, and have a low reproductive rate. Age estimates extend up to 100 yr, although 60-80 yr is considered a more reasonable estimate. Age to sexual maturity ranges from 10 to 20 yr, with possible latitudinal association (Alford 1980, Iverson 1980, Landers et al. 1982, Wright 1982, Doonan 1986, McLaughlin 1990, Mushinsky et al. 1994). Gopher tortoises lay one clutch of eggs annually, generally ranging from 2-10 eggs in size, with the average being 4-8 (Dietlein and Franz 1979, Iverson 1980, Landers et al. 1980, Linley and Mushinsky 1994). In some areas with excellent food resources and large tortoises, clutch sizes may average 9-10, with a range of 6-14 eggs (McLaughlin, 1990). Although some wild individuals have produced over 20 eggs in one clutch (Godley 1989, L. Macdonald, personal communication), this is very unusual. With predation rates of up to 95% on eggs and hatchlings, and further high losses of juveniles aged 1-5 yr, less than 10%, and possibly as low as 1%, of eggs laid eventually produce reproductive adults (Douglass and Winegarner 1977, Auffenberg and Iverson 1979, Alford 1980, Iverson 1980, Landers et al. 1982, Diemer 1986, Wilson 1991). The gopher tortoise is listed in Appendix II of the Convention on International Trade in Endangered Species of Wild Fauna and Flora (C.I.T.E.S.), which requires permits for the exportation of the species from the U. S. to any signatory nation, or for reexportation (Levell 1995). However, if federal or state regulations are more restrictive

PAGE 13

than C.I.T.E.S., those take precedence. Legal protection is extended to the species in all states within the range, although the levels of protection vary. The populations west of the Tombigbee and Mobile Rivers in Alabama, Mississippi and Louisiana are on the federal threatened species list, which prohibits the taking, exportation, or interstate movement of individuals originating from that region without permit, and the possession, transportation, purchase, and/or sale of illegally obtained specimens. Permits issued by the U. S. Department of Interior can be obtained for scientific research on wild populations, and possession of limited numbers of individuals for exhibition, education, and/or research. State permits are required also. The gopher tortoise is listed in Alabama as a protected nongame species, in Mississippi as endangered, and in Louisiana as threatened. Louisiana regulations also prohibit the use of gasoline, chemicals, or volatile substances to flush reptiles from burrows or other hiding places. Georgia lists the species as threatened and issues scientific collection permits only to qualified institutions and individuals for educational or research purposes. Although the law states that burrows may not be disturbed nor destroyed, nor may explosives, chemicals or smoke be introduced into them to drive out wildlife, the code explicitly exempts poisonous snakes. Because venomous snakes, particularly rattlesnakes, use gopher tortoise burrows, tortoises can be adversely impacted by such activities. South Carolina was the last state to extend legal protection to gopher tortoises (Mann 1990, Levell 1995), and now lists the species as endangered, with permits required for any activities involving tortoises. In Florida, where the gopher tortoise is listed as a species of special concern (Wood 1996), regulations prohibit taking and disturbing of tortoises and their habitats, although exceptions are granted regularly for development, agriculture, and mining operations. In the past, human consumption of

PAGE 14

tortoises was a major impact on regional distribution, and current poaching activities may extirpate local aggregations (Diemer 1989, Mann 1990). Large scale conversion of longleaf pine habitats to slash pine plantations, transformation of native pasture or savanna to "improved" pasture, citrus groves, or row crops, mining operations such as phosphate, mineral sand, and gravel mines, and urban/suburban development are the main threats to continued gopher tortoise survival today (Diemer 1986, Cox et al. 1987, Diemer and Moore 1994). Lack of natural fire regimes due to suppression efforts by humans may alter vegetation mosaics in rermining habitat, rendering them less suitable to continued maintenance of tortoise populations (Mushinsky 1986, Mushinsky and Gibson 1991). Although tortoise numbers increase with increasing areal extent of available habitat, densities remain constant or decrease (Mushinsky and McCoy 1994). Fragmentation of mainland areas, with ever smaller islands of suitable habitat surrounded by agricultural and urban development, may force tortoises into higher density populations than would occur normally. Vegetational changes resulting from reduced fire incidence in small areas, particularly increased canopy closure and decreased herbaceous vegetation (Mushinsky 1985, 1986), can lead to decreases in reproductive rates and juvenile survival (Auffenberg and Franz 1982, McLaughlin 1990, Mushinsky and McCoy 1994). Increased intraspecific interactions at higher densities (McRae et al. 1981) may lead to elevated physiological stress, potentially affecting immune system function and rendering animals more susceptible to disease. Increased densities and number of interactions multiply the opportunities for transmission of communicable agents, making fragmented populations more likely to sustain high morbidity during epizootics. If mortality is high, lowered reproduction due to habitat degradation may be insufficient for population recovery.

PAGE 15

6 Along with increased physiological stress placed on tortoise populations from human activities, there may be toxicological or immunological stress from chemicals introduced into the environment. No research has been conducted into the effects of herbicides, fungicides, insecticides, and fertilizers on gopher tortoise health, growth, or reproduction. Investigations of disease in free-ranging populations of tortoises have begun only recently. Tortoises and Upper Respiratory Tract Disease Although individual captive and wild gopher tortoises have been observed with clinical signs of respiratory diseases for over 20 yr (E. R. Jacobson, unpublished data), the first documentation of a larger-scale disease outbreak was in 1989, when an epizootic of upper respiratory tract disease (URTD) was documented on Sanibel Island, Lee County, Florida (G. S. McLaughlin and M. Elie, unpub. data). With the loss of 25-50% of breeding-age adults in one population, recovery could take 50-150 yr (G. S. McLaughlin unpub. data), barring further major losses and without substantial habitat improvement leading to increased recruitment. In the 1980s, large-scale population reductions (33-76% over 10 yr) of desert tortoises (Gopherus agassizii) were documented at several sites in the western Mojave Desert of California and at one site in the eastern Mojave (Corn 1994, Berry in press). Tortoises with clinical signs of URTD were observed among the remaining populations at several sites (Knowles 1989; Berry 1990, in press). As a result of the declines, tortoises in the Mojave Desert north and west of the Colorado River were declared threatened (U.S. Fish and Wildlife Service 1990).

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7 Clinical signs of URTD in gopher and desert tortoises include serous, mucoid, or purulent discharge from the nares, excessive tearing to purulent ocular discharge, conjunctivitis, and edema of the eyelids and ocular glands (Jacobson et al. 1991, G. S. McLaughlin personal observations). Individual infected tortoises vary in the suite of signs they have, and the severity can vary from day to day. Nares may become occluded with caseous exudate, preventing externally visible nasal discharge. Lymphocytic infiltration of the corneas, while rare, may decrease an animal's ability to forage or avoid predators. Tortoises may become lethargic and anorectic, leading to dehydration, emaciation, and eventual death from cachexia. Lethargy, nonresponsiveness to stimuli, and altered behavior patterns—such as basking at lower temperatures than normal— may render a tortoise more susceptible to predation. Moribund animals often develop petechial to ecchymotic hemorrhages under the scutes, especially visible on the plastron (G. S. McLaughlin pers. obs.), which may be due to septicemia caused by secondary infection with opportunistic bacteria. Several agents were hypothesized to cause respiratory tract disease in tortoises, including viruses, Mycoplasma sp. (Lawrence and Needham 1985), and Pasteurella testudinis (Snipes and Biberstein 1982). However, experimental infections to determine the etiologic agent were not conducted. Beginning in 1989, efforts were undertaken to determine the etiology of URTD. Clinically ill and healthy desert tortoises from California were examined and samples collected, either from live animals or at necropsy. Hematologic and serum biochemical evaluations, liver vitamin and metal determinations, and pathologic and microbial investigations were conducted (Jacobson et al. 1991).

PAGE 17

8 Hemoglobin concentration and phosphorous levels were lower in ill than in healthy tortoises, and ill tortoises had higher sodium, blood urea nitrogen (BUN), creatinine, serum glutamic oxalacetic transaminase (SGOT) activity, and total cholesterol levels. Levels of liver and serum vitamins A and E did not differ between healthy and ill tortoises, nor did hver values of selenium, copper, lead, or cadmium. However, iron and mercury were higher in ill than in healthy tortoises. Healthy tortoises did not exhibit any histological abnormalities in nasal mucosa or other tissues. Ill tortoises had less fat than controls, atrophied thymuses, increased numbers of lymphocytes in the sinusoids of the spleens, and increased amounts of iron granules in the hepatocytes. The most striking and consistent changes were in the nasal passage and cavity tissues (Jacobson et al. 1991). Grossly, passages and cavities of ill tortoises contained moderate to large amounts of serous to purulent exudate. Histologically, the tissues exhibited loss of mucosal glands and goblet cells, proliferation of epithelial cells, infiltrates of lymphocytes and histiocytes, loss of cilia, dysplasia of the olfactory epithelia with infiltration of heterophils, basal cell hyperplasia, occasional squamous metaplasia, and occasional erosion and ulceration of mucosal epithelia. By electron microscopy, pleomorphic organisms resembling Mycoplasma sp. were seen on cell surfaces and tightly adhered to cell membranes of ill tortoises, but not healthy ones. Significantly more Pasteurella testudinis were isolated from ill than healthy tortoises, and Bacillus sp. and Mycoplasma-hkQ organisms were isolated only from ill tortoises. By electron microscopy, the latter resembled those seen on the nasal cavity tissues of ill tortoises (Jacobson et al. 1991). The mycoplasma was determined to be a new species, provisionally named Mycoplasma agassizii (Brown et al. 1994). An enzyme-linked immunosorbent assay

PAGE 18

9 (ELISA) to detect antibodies against the mycoplasma in plasma and serum samples was developed (Schumacher et al. 1993), and experiments were undertaken to fulfill Koch's postulates. The disease was induced by inoculation of tortoises with pure cultures of the mycoplasma, but not Pasteurella testudinis (Brown et al. 1994). Histologically, the lesions were consistent with those seen in the previously examined naturally infected tortoises. Additional work led to the development of a polymerase chain reaction (PCR) test to detect the bacteria in nasal flush and swab samples (Brown et al. 1995). Histologic examination of nasal tissues and microbiologic evaluation of flush and swab samples from clinically healthy and clinically ill tortoises from Las Vegas Valley, Nevada, resulted in findings in the ill tortoises similar to those in the California tortoises, with 92% showing lesions of URTD, 50% being culture positive for M. agassizii, and 100% reacting positively in the ELISA (Jacobson et al. 1995). However, 73% of the "healthy" tortoises had lesions consistent with URTD, 50% were culture positive forM agassizii, and 42% were seropositive for antibodies against the bacteria. These findings demonstrate that the disease can exist in a subclinical form in a substantial proportion of a population. An annual cycle of convalescence and recrudescence of clinical signs has been seen in captive desert (I. M. Schumacher, pers. comm.) and gopher (D. L. Morris, pers. comm.) tortoises. Other mycoplasmal diseases also can exist as chronic, subclinical infections, with recurrence of clinical signs and increases in transmission potential when the host is stressed (see below). Annual fluctuations in temperature, rainfall, and forage availability may be sufficient to cause detectable outbreaks in an infected population. Increased morbidity and mortality may occur in times of unusually severe environmental stress, such as prolonged drought, hurricanes, excessive rainfall with flooding of burrows,

PAGE 19

or very cold winters. Human impacts on tortoises and their habitat, whether through disruption of normal behavior patterns, degradation of habitat through agriculture, silviculture, rnining or development operations, or pollution, may cause sufficient physiological stress to trigger proliferation of the mycoplasma and recurrence of signs. Capturing and transporting of tortoises during relocation, restocking and repatriation efforts also may be significant sources of stress. Mycoplasmal Respiratory Diseases in Domestic Animals and Humans Mycoplasmas cause respiratory disease in other taxa, including rodents, poultry, swine, ruminants, and humans. All have similar pathological effects, which are described below, and many are exacerbated by concurrent infections or environmental stress. Murine respiratory disease caused by Mycoplasma pulmonis has caused problems in laboratory settings for more than 70 yr (Lindsey et al. 1971). It has necessitated complex and expensive control measures, including various types of isolation mechanisms and cesarean delivery of mice and rat pups to reduce the prevalence of M. pulmonis infections in colonies (Cassell et al. 1984, Davis et al. 1987). Interactions among environment, host, and strain factors influence the impacts at individual and population levels (Simecka et al. 1992). Histologically, lesions are characterized by lymphoid hyperplasia and chronic inflammation (Cassell et al. 1985). Poultry respiratory diseases can be caused by viruses, mycoplasmas, and other bacteria, singly or in combination. Without concurrent viral or other bacterial infections, disease can be mild or subclinical (Kerr and Olson 1967). Environmental factors, particularly dust and ammonia levels, as well as strain differences, affect the severity of

PAGE 20

11 outbreaks of mycoplasmosis. Other stress factors, such as crowding and cold weather, also influence morbidity and mortality (Jordan 1972). Mycoplasma gallisepticum causes chronic respiratory disease in chickens and sinusitis in turkeys, and affects ring-necked pheasants (Phasiantis colchicus), chukar (Alectoris chukar), northern bobwhite (Colinus virginianus), common peafowl (Pavo cristatus) (Yoder 1991) and Japanese quail {Coturnix japonica) (do Nascimento and do Nascimento 1986). Lower respiratory tract complications are seen rarely in turkeys (Simecka et al. 1992). Histologically, hypertrophy and hyperplasia of respiratory epithelia, including mucous cells, are seen, as is lymphoid infiltration of the submucosa. Heterophilic exudate is seen in the airways (Nunoya et al. 1987, Trampel and Fletcher 1981). Mycoplasma meleagridis infection is seen primarily in chicks and poults up to 10 wks of age, and is sexually transmitted. Turkeys, Japanese quaiL and peafowl develop air sacculitis, sometimes with accompanying tracheitis and pneumonia, but not sinusitis. Histologically, the lesions are characterized by hypertrophy and hyperplasia of the air sac epithelia, with edema and lymphoid infiltration (Stipkovits 1979). Young chickens and turkeys also are susceptible to M. synoviae, usually in conjunction with Newcastle disease virus or infectious bronchitis (Hopkins and Yoder 1984, Springer et al. 1974). As with M. gallisepticum, turkeys develop sinusitis (Stipkovits 1979). All three bacteria can be transmitted vertically (i.e., via the egg) (Simecka et al. 1992). Swine develop mild pneumonia when infected by M. hyopneumoniae, and although mortality is virtually nil, the disease is chronic, causing slow growth and reduced weight gains, decreasing profitability (Oboegbulem 1981, Jericho 1986). Although signs can disappear, recurrences will occur with weather changes, viral infections, and other

PAGE 21

stressors (Whittlestone 1976). Lesions are alveolar, initially characterized by neutrophilic infiltrates and later by lymphocytes and macrophages (Baskerville 1972). Contagious bovine pleuropneumonia (CBPP), although not present in the United States, is an economically important disease of cattle and water buffalo caused by M. mycoides subsp. mycoides (Howard and Gourlay 1978, Trichard et al. 1989). Morbidity rates can reach 100% with up to 50% mortality, and recovered animals maintain a carrier state. Neutrophilic exudate in the airways, serous exudate in the pleural cavity, serofibrinous exudate in alveoli, and edema and necrosis of regional lymph nodes and interlobular septa are seen at necropsy. Interseptal, peribronchial, and perivascular lymphoid infiltration can be seen histologically (Hudson 1971). Calf pneumonia, with high mortality among dairy calves, is caused primarily by viruses, but can be caused by either M bovis or M dispar (Stalheim 1983, Bryson 1985). Infection with the former is characterized by peribronchiolar and alveolar monocyte infiltration, while the latter causes interstitial pneumonia with monocyte infiltration of alveolar walls, but not peribronchiolar infiltration. Mycoplasma dispar also causes superficial and asymptomatic infection of respiratory mucosa (Woldehiwet et al. 1990). Lambs are susceptible to infection with M. ovipneumoniae, which is transmitted to neonates from the ewes. Pneumonia, characterized by coughing, sneezing, nasal discharge, fatigue, and poor weight gain, can develop as colostral antibodies wane. Lesions are characterized by alveolar proliferation, nodular lymphoid hyperplasia, and peribronchial and perivascular lymphoid infiltration (Carmichael et al. 1972, Foggie et al. 1976). Severity can be exasperated by infection with Pasteurella haemolytica biotype A (Jones etal. 1982).

PAGE 22

13 Goats can suffer 60-100% mortality when infected by M. mycoides subsp. mycoides, M. mycoides capri, or Mycoplasma strain F38, making contagious caprine pleuropneumonia the most economically important goat disease. Pathologically, signs and lesions are similar to those seen in CBPP, with the addition of polyarthritis (Cottew 1984). Humans are susceptible to infection with M. pneumoniae, which causes tracheobronchitis and, less commonly, primary atypical or walking, pneumonia (Chanock et al. 1963, Clyde 1983). The upper respiratory tract is affected, not the alveoli, and otitis media is seen also (Clyde 1979, Mansel et al. 1989). Immunopathological sequelae, although rare, can include arthritis, dermal lesions, cardiopathy, and neurological complications (Johnston et al. 1983, Lyell et al. 1967, Naftalin et al. 1974, Murray et al. 1975). Histologically, perivascular and peribronchial monocytic infiltrations with some neutrophilic exudate are seen (Dajani et al. 1965). Most commonly affected are 5-9 yr old children, with a decline in incidence until about age 25, then an increase in 30-40 yr olds (Denny et al. 1971). The disease is endemic, with slight increases in late summer and fall, and cyclic epidemics occurring at 4-7 yr intervals (Krause and Taylor-Robinson 1992). Mycoplasmal Diseases of Wildlife Research on the impact of mycoplasmosis on wildlife is limited, but recent developments are provoking interest. In early 1994, house finches (Carpodacus mexicanus) in the mid-Atlantic and northeastern regions of the United States were found with conjunctivitis, rhinitis, and sinusitis. Many birds were found dead or dying and were submitted to wildlife care facilities, veterinary hospitals, and state agricultural laboratories. Many of those birds had lesions that were histologically compatible with mycoplasma

PAGE 23

14 infections, and were infected with M. gallisepticum (Ley et al. 1996, Luttrell et al. 1996). Since then, the organism has been associated with morbidity and mortality in other species of passerines, including goldfinches (Carduelis tristis) (Nettles 1996), and was transmitted to a blue jay (Cyannocitta cristata) in a rehabilitation facility (Ley et al. 1996). Conjunctivitis and M. gallisepticum also were found in house finches in Georgia (Luttrell et al. 1996). The potential for this organism to spread over a large area due to the long distances traveled by migratory birds, the mixed species flocks in which they congregate, and local concentrations of many species around bird feeders is of great concern to many ornithologists and ecologists, as well as poultry producers. There are indications that the strain, while highly pathogenic to chickens, is not readily transmissible to poultry under natural conditions (Nettles 1996). A new species of mycoplasma (M sturni) was isolated from the conjunctiva of a European starling (Sturna vulgaris) with conjunctivitis found in Connecticut during the epornitic (Forsyth et al. 1996). Although it was isolated in pure culture, it was described as growing rapidly, whereas pathogenic mycoplasmas typically grow slowly; therefore, it could have overgrown M. gallisepticum if that species had been present. The pathogenicity of M. sturni needs to be investigated. Three new species of mycoplasmas have recently been described from raptors in Europe (Poveda et al. 1994). All were associated with respiratory diseases clinically and histologically consistent with those caused by Mycoplasma spp., with lesions including hyperplasia of mucous glands, lymphoid hyperplasia, and perivascular cuffing (Poveda et al. 1990). The species were isolated from buzzards (Buteo buteo), saker falcons (Falco cherrug), and griffon vultures (Gyps fulvus), and have been named, respectively, M buteonis, M. falconis, and M. gypis. Pathogenicity of these species, their distribution in

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15 wild, free-ranging birds, and their potential impacts on the populations need to be further investigated. Several species of mycoplasmas, including M cloacale (Bradbury et al. 1987, Goldberg et al. 1995) and M anatis (Ivanics et al. 1988, Poveda et al. 1990, Goldberg et al. 1995), have been isolated from semi-domestic and wild ducks and other avian species throughout the world. Stipkovits et al. (1986) reported isolation of M. cloacale from geese with inflammation of the cloaca and phallus, but Goldberg et al. (1995) found no association of M. cloacale with disease in wild mallards (Anas platyrhynchos), black ducks (A. rubripes), or canvasbacks (A. valisneria). Stipkovits (1979) reported pathogenicity of M. anatis to domestic ducklings and eggs, and neurological signs have been recorded in ducks infected with M. anatis (Ivanics et al. 1988). Samuel et al. (1995) infected game-farm mallard eggs with M. anatis and found reduced hatchling success, hatchling size and growth rates. Hatchlings infected at 1 d of age did not have slower growth rates. Goldberg et al. (1995) found esophagitis, tracheitis, and vaginitis in female mallards from which M. anatis was isolated, and presented evidence for vertical transmission of M. anatis in a wild gadwall (Anas strepera). Potentially, M. anatis infections could reduce recruitment in wild duck populations. Experimental infection of ducklings with M. gallisepticum resulted in suppressed growth rates (Stipkovits 1979). Four other unidentified mycoplasmas were isolated from mallards, gadwalls, and black ducks, but could not be associated definitively with disease (Goldberg et al. 1995). Although M. gallisepticum has been isolated from wild turkeys, most cases have occurred in birds with close association to domestic poultry (Davidson et al. 1982, Jessup et al. 1983, Luttrell et al. 1991, Fritz et al. 1992). Even though experimental infection of

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16 pen-reared wild turkeys has resulted in decreased productivity (Rocke et al. 1988), there is little evidence to indicate that infection with M. gallisepticum strains commonly occurring in domestic fowl poses a threat to wild turkey populations (Luttrell et al. 1991), or that wild turkey populations are important in the epizootiology of M gallisepticum (Davidson et al. 1988). Mycoplasma gallopavonis has been isolated from wild turkeys in Texas (Rocke and Yuill 1987), South Carolina, Georgia (Luttrell et al. 1991), Colorado, New Mexico, and Oklahoma (Fritz et al. 1992). Although lethal to experimentally infected domestic turkey eggs (Rocke and Yuill 1987), the pathogenicity of M. gallopavonis to wild turkeys has not been investigated. Mycoplasma synoviae and other, untyped, Mycoplasma spp. were isolated from turkeys in Arizona (Fritz et al. 1992), but no association with disease was found. As with M. gallisepticum, wild turkeys do not appear to be important in the epizootiology of M. synoviae or M. meleagridis (Davidson et al. 1988). In 1993 an epizootic of polyarthritis occurred in juvenile farmed crocodiles (Crocodylus niloticus) in Zimbabwe. The outbreak was characterized by high morbidity, but low mortality. A mycoplasma was isolated, determined to be a previously unrecognized species, and named M. crocodyli (Mohan et. al 1995). In 1995 a die-off of captive adult American alligators (Alligator mississippiensis) at a private facility in Florida, with very high mortality, was associated with systemic infection with a different species of mycoplasma, also previously unrecognized (Brown et al. 1996a). The name M. lacerti has been proposed. In addition to M. agassizii, a second mycoplasma was found in desert tortoises with evidence of URTD, including clinical signs, histologic lesions, and/or positive ELISA tests (Brown et al. 1995). In a small pilot study, that organism was found

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17 to cause disease and seroconversion in gopher tortoises previously unexposed to either mycoplasma (D. R. Brown, et al., unpub data). Mycoplasmas have been isolated from bighorn sheep (Ovis canadensis) with pneumonia associated with P. haemolytica, and there is concern that these may have a role in the disease process (D. Hunter, pers. comm.). A mycoplasma isolated from dead and live captive bighorn sheep with pneumonia was typed asM arginini (Woolf et al. 1970). Apparently, no further work has been done regarding the pathogenicity of that strain to bighorn. With continued exposure of bighorn to domestic livestock, Mycoplasma spp. could interfere with recovery efforts. An outbreak of pneumonia associated with M. ovipneumoniae infection occurred in captive Dall's sheep (Ovis dalli dalli) following indirect exposure to domestic sheep (Black et al. 1988). Although Dall's sheep are unlikely to be exposed to domestic livestock in their native habitat, it is clear that captive herds or those in transit must be protected from exposure to pathogens. Behymer et al. ( 1989) detected antibodies to Mycoplasma spp. in mule deer (Odocoileus hemionus) in California, but there was no association with disease, and isolations were not attempted. One-humped camels (Camelus dromedarius) and African buffalo (Syncerus caffer) on a game farm in Kenya had antibodies against Mycoplasma strain F38, and camels were seropositive for M. mycoides mycoides, but no isolations were made and no disease was seen (Paling et al. 1988). Kirchoff et al. (1996) found, in a survey of captive arthritic elephants (Elephas maximus and Loxodonta africana), that about 60% of the females harbored a new mycoplasma in the urogenital tract, although none was recovered from males. It is not known if the mycoplasma, named M. elephantis, causes arthritis.

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18 Chronic Manifestations of Mycoplasmal Infections Many mycoplasmal diseases are characterized by an overaggressive or inappropriate immune response by the host, eventually leading to autoimmune damage to the affected sites, whether respiratory or urogenital tract, joints, heart, skin, or other organ systems (Krause and TaylorRobinson 1992, Simecka et al. 1992, Cole 1996). Infected hosts may be more susceptible to secondary infections with other bacteria or viruses. In wildlife species, such complications may reduce the fitness of the animals by altering behavior, leading to decreased foraging efficiency, increased susceptibility to predators, or diminished mate seeking behavior. Energy that normally would be allocated to reproduction may be needed to repair or compensate for damage to multiple organ systems. Therefore, even if mycoplasmal infections do not cause mortality directly, they can affect individual and population viability. Project Overview and Specific Objectives Due to the listing by the Florida Game and Fresh Water Fish Commission of the gopher tortoise as a species of special concern, and the subsequent permitting of over 450 relocations involving more than 8000 tortoises, particular attention has been focused on the dynamics and persistence of both natural and relocated populations. Understanding the effects of URTD on individuals and populations is essential for proper management of rermining populations; therefore, a study was begun in 1993 on the etiology, pathology, and diagnosis of URTD in gopher tortoises. The original objectives were 1) to describe the pathology of natural infections and identify possible etiologic agents, 2) to perform

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19 experimental infections to determine conclusively the etiologic agent and dose-related effects, 3) to refine the PCR test and delineate further the taxonomy of the tortoise mycoplasmas, 4) to characterize the immune responses of gopher tortoises to URTD pathogens, and 5) to compare the efficacy of culture, PCR and serology in detecting new infections. As the original experiments were conducted and concern over the continued wellbeing of tortoise populations throughout the range increased, additional questions were raised. Those were concerned primarily with management and conservation of gopher tortoises, particularly with respect to relocation, repatriation, and restocking efforts. Several questions were raised repeatedly in meetings with state and federal agency personnel, particularly regarding possible immunity to the organism and factors related to transmission. In order to address those questions additional objectives were established, as delineated in specific objectives 2-7 below, and discussed in Chapters 3 through 8. Chapter 2 will detail the methods common to fulfilling the objectives. This dissertation addresses the following specific objectives: 1) To describe the clinical presentation and lesions of natural infections by M agassizii in gopher tortoises and compare the lesions to those found in desert tortoises (Chapter 3). 2) To test the hypothesis that tortoises that have produced antibodies against M. agassizii are protected against reinfection with the organism and subsequent development of URTD (Chapter 4). 3) To assess the probability of horizontal transmission of M agassizii between adult tortoises and to test the hypothesis that transmission is more likely to occur from an

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20 infected host that is clinically ill and PCR or culture positive than from a seropositive host that is not clinically ill or is clinically ill but culture or PCR negative (Chapter 5). 4) To determine if M. agassizii is transmitted vertically (Chapter 6). 5) To assess the relationship of antibodies in eggs and hatchling serum to those in maternal serum (Chapter 6). 6) To collect preliminary data to test the hypothesis that M. agassizii can be transmitted environmentally (e.g., in burrows) (Chapter 7). 7) To discuss the implications of the findings of the above research and additional concurrent research on conservation and management of gopher and other tortoises (Chapter 8).

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CHAPTER 2 METHODS Tortoises, Intake Procedures. Clinical Assessments and Sampling Methods For the natural infection studies, tortoises were obtained from various locations in the state of Florida under Florida Game and Fresh Water Fish Commission permits number WX93227 issued to Elliott R. Jacobson and number WX94037 issued to Mary B. Brown. Tortoises were processed the day of arrival at the University of Florida (UF), Gainesville. For the natural and experimental infection studies, gopher tortoises were transferred from a development site in central Florida to UF in April, July, and August 1994 and April 1995, and processed the day following arrival. Tortoises were examined for clinical signs of URTD: nasal and ocular discharge, palpebral edema, and conjunctivitis. The signs were graded individually on a scale of 0-3, none to severe. Total clinical sign score was calculated as the nasal discharge score plus the mean of three ocular sign scores (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 a(iministered at 20 mg/kg. Straight line carapace length, thickness, and width were measured to the nearest cm with forestry calipers. 21

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22 A 2-3 ml blood sample was drawn from the jugular or brachial vein and placed in a lithium heparin Vacutainer tube (Becton Dickinson and Company, Rutherford, New Jersey). Blood was centrifuged and an aliquot of plasma was removed for antibody screening by an enzyme-linked immunosorbent assay (ELISA) (see below). After cleansing the area around the nares with alcohol dampened gauze, nasal flush samples were collected by flushing with approximately 0.5 ml sterile SP4 broth using a 1 ml syringe without needle. Calcium alginate-tipped swabs were inserted gently into the nares, and a sample was obtained and streaked onto SP4 agar plates. Culture Procedures A 100 pi aliquot of the flush sample was used for polymerase chain reaction (PCR) analysis; the remaining sample was serially diluted tenfold to 10" 2 and incubated at 30°C for a maximum of three weeks, or until determined to be positive or contaminated. Twenty pi of each dilution were placed on SP4 agar and incubated at 30°C and 5% CO, as were the streak plates. Plates were examined regularly for a maximum of six weeks to detect the growth of mycoplasma. In the second year of the study, broth cultures were incubated for 24-48 hr before removing the aliquot for PCR, as that modification increased the sensitivity of the PCR (G. S. McLaughlin and D. R Brown, unpub. data). PCR Procedure Nasal aspirate samples were analyzed for the presence of Mycoplasma agassizii DNA based upon PCR amplification of the 16S rRNA gene (Brown et al. 1995). Nasal flush and culture samples were centrifuged for 60 min at 14,000 rpm at 4°C, and the

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23 supernatant aspirated. Three to four microliters of 20 ug/ml proteinase K (Sigma, St. Louis, MO) in 20 pi lysis buffer ( 100 mM tris pH 7.5, 6.5 mM DTT, 0.05% Tween 20) were added to the pellets, which were resuspended, and the samples were incubated at 37°C for 8-16 hours. After denaturing the proteinase K at 97°C for 15 min, 5 pi of each sample were removed and added to 45 pi of reaction solution containing two primers for the 16S rRNA gene at 1 pM each, deoxynucleoside triphosphates at 200 pM, 2.0 mM MgCL and 2.5 units of Taq polymerase (Promega, Madison, WI). 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 ', Brown et al. 1995) and a Mycoplasma genus-specific region [anti-sense strand nt 1055 to 1031, 5'-TGCACCATCTGTCACTCTGTTAACCTC-3', Van Kuppeveld 1992]. Samples were subjected to 50 cycles of template denaturation for 45 sec at 94°C, primer annealing for 1 min at 55°C, and polymerization for 45 sec at 72°C, followed by 10 min at 72°C. Positive samples yielded 576 base pair (bp) products that were visualized by combining 15 pi of product with 2 pi bromphenol blue in 50% glycerol solution and electrophoresing on etWdium bromide-stained 1.5% agarose gels in tris-borate-EDTA buffer. Positive control samples using 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, Hind III digest of phage DNA, was included on each gel. In order to confirm that the isolates obtained from naturally and experimentally infected tortoises were M. agassizii, an additional procedure, restriction fragment length polymorphism (RFLP) analysis, was conducted on at least one isolate from each tortoise. Twenty microliter samples of products from the above amplification procedure were

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24 incubated with 10-20 units of the endonuclease Agel (New England Biolabs, Inc., Beverly, MA), which cuts the M. agassizii amplification product at nt 613, and 5 pi of reaction buffer, at 25°C for one hour, and the products electrophoresed as above. The procedure resulted in products of 434 and 142 bp fromM agassizii-positivQ samples, and no change in non-M agassizii-s&mples. ELISA Procedure An aliquot of plasma from each sample was used for determinations of levels of antibodies specific for M. agassizii (Schumacher et al. 1993). Ninetysixwell microtiter plates (Maxisorp F96, Nunc, Kamstrup, Denmark) were coated with 50 pi of a whole-cell lysate ofM agassizii strain 723 at lOpg/ml in phosphate buffered saline with 0.02% azide (PBS-AZ). 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 ul/well PBST containing 5% non-fat dry milk (PBS-TM). Following washing, 50 pi of plasma diluted appropriately for the specific study with PBS-TM were added to individual wells in duplicate or triplicate, and the plates were incubated at room temperature for 60 min. The plates were washed, 50 pl/well of a biotinylated monoclonal antibody (MAb HL673) against the light chain of desert tortoise immunoglobulins IgY and IgM at 1 pg/ml in PBS-TM was added, and plates were incubated for 60 min. Following washing, a conjugate of alkaline phosphatase and streptavidin (AP-S; Zymed Laboratories, Inc., San Francisco, CA) at 1:2000 in PBSAZ was added at 50 pl/welL plates were incubated for 60 min, and washed. Substrate, pnitrophenyl phosphate disodium (pNPP; Sigma), was prepared at 1 mg/ml in 0.01 M

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25 sodium bicarbonate, pH 9.6, with 2 mM MgCl 2 , and added to wells at 100 ul/well. Plates were incubated for 60 min in the dark, then read at 405 nm on a microplate reader (EAR 400 AT, SLT Labinstruments, Salzburg, Austria). The mean of two or three wells coated with antigen and incubated with conjugate and substrate only was used as the blank. A positive control, plasma from a naturally infected gopher tortoise from Sanibel Island, and a negative control, plasma from an uninfected tortoise from Orange County, were included on each plate. Results from the ELISA were optical density (OD) readings from the microplate reader. The OD readings reflected the intensity of the yellow color developed when all components of the reaction (specific tortoise antibodies against M. agassizii, biotinylated MAb HL673, AP-S, and /?NPP) were present. The readings are on a continuous scale, but were interpreted categorically by calculating the ratio of the sample readings to the negative control reading. Ratio values less than or equal to 2.0 were considered negative, those greater than 2.0 and less than or equal to 3.0 were classified as suspect, and those greater than 3.0 were classified as positive. Study Group Assignment Tortoises exhibiting one or more signs of disease, or with a positive culture or PCR result, were designated as diseased. Animals were designated clinically healthy if found free of signs of URTD, and with negative culture and PCR results. Tortoises were assigned to study groups based on sex, weight, clinical assessments, culture, PCR and ELISA results, and placed in the appropriate pens.

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26 Husbandry Procedures Tortoises were housed singly or in pairs in outdoor pens at the UF Animal Resource Farm (ARF). There were four groups of 10 pens in a larger, chain linkfenced enclosure (Fig. 2-1). Pens were approximately 21 m 2 , constructed of a wooden frame with sheet metal extending vertically approximately 0.7 m above and below ground, partially covered by shade cloth, and provided with an artificial burrow, a water dish, and a cement feeding stone. Pens were observed daily by ARF staff, and watered daily in the summer and as needed in the winter. The tortoises were fed a salad of mixed vegetables three times per wk, and fruit was provided on an occasional basis. In addition, I observed the pens and tortoises from three to seven days each week, with some days including multiple observations. Because of individual behavior patterns, not every tortoise was observed at each time point. The amount of food eaten was recorded for each pen the day following feeding, and the stones were cleaned. Remaining food was collected and bagged separately for each section, using brushes and dust pans assigned to that section. Stones were then sprayed with a dilute (1:30) bleach solution, allowed to soak for a few minutes, and hosed off. Water dishes were rinsed and filled daily, and bleached and scrubbed as necessary. 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 a different tortoise. Used gloves were placed in plastic garbage bags and disposed of properly. Before entering the first pen on a given day, personnel sprayed their footwear with a dilute bleach solution. Footwear was sprayed with bleach before leaving

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27 a pen. Animals were captured by hand or using wire cage-type traps (Tomahawk Live Trap Company, Tomahawk, WI) that were covered with brown paper to protect the animals from the weather. Traps were cleaned, sprayed with bleach solution, and allowed to air dry following each use. 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, WI) for transport and holding. Containers were bleached, scrubbed, and washed in an automatic cage washer before reuse. D C A B Figure 2-1. Layout of tortoise pens at the University of Florida Animal Resource Farm The outer fence was chain link, and the inner fence and pen dividers were corrugated sheet metal on a wooden frame. Necropsy Procedures All diseased and selected healthy tortoises were euthanatized with a combination of drugs. Ketamine was administered intramuscularly at 60 80 mg/kg followed by a

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28 concentrated barbiturate solution (Socumb, The Butler Company, Columbus, Ohio, USA) intraperitoneally at 1 ml/kg. Once the tortoises showed complete muscle relaxation and were unresponsive to painful stimulation, they were exsanguinated via a 23 gauge butterfly catheter inserted into the carotid artery and then decapitated. Flush and swab samples were collected as previously described, then the head was bisected longitudinally with an electric saw. Following bisection, the cartilage over each nasal cavity was reflected aseptically, and flushes and swabs of 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 and the following tissues were collected, fixed in neutral buffered 10% formalin, sectioned at 5-6 um, and stained with hematoxylin and eosin: glottis, cranial trachea, tracheal bifurcation, left lung, right lung, thyroid, heart, brain, thymus, esophagus, stomach, small intestine, pancreas, large intestine, cloaca, spleen, liver, left and right kidney, bladder, right and left gonads, chin gland, buccal salivary gland, chin gland, and tongue. Tissues were examined by light microscopy and abnormalities or changes were recorded. Histopathology Procedures For histopathologic studies, heads were fixed in 10% neutral buffered formalin (NBF), decalcified, embedded in paraffin, sectioned longitudinally at 5-6 um, and stained with hematoxylin and eosin. Sections were examined by light microscopy and classified on a scale of 0 to 5, with 0 being normal and 5 exhibiting severe inflammation and / or changes. Changes in the epithelium and submucosa were recorded separately.

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29 The following criteria were utilized for grading lesions: Normal (score = 0): Occasional small subepithelial lymphoid aggregates; rare heterophils in the lamina propria. No changes in mucosal or glandular epithelium; no edema. Mild (1): Multifocal small subepithelial lymphoid aggregates; multifocally, small numbers of heterophils, lymphocytes, and plasma cells in the lamina propria; mild edema in lamina propria; minimal changes in mucosal epithelium. Moderate (2-3): 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; proliferation and disorganization of the basal epithelium. Severe (4-5): Focally extensive to diffiise band of lymphocytes and plasma cells subjacent to and obscuring the overlying mucosal epithelium; large numbers of heterophils in lamina propria and infiltrating 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; occasional squamous metaplasia. Statistical Analyses Statistical analyses were performed using SAS (SAS Institute, 1988) or SigmaStat for Windows, Version 1.0 (Fox et al. 1994). Because the analyses varied for each experiment, the specific methods will be addressed in the appropriate chapters.

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CHAPTER 3 NATURALLY OCCURRING UPPER RESPIRATORY TRACT DISEASE Methods Tortoises Twenty-three gopher tortoises from the following locations in Florida (Figure 3-1) were transported to UF from August 1993 to September 1995: Alachua County (n = 2), Sanibel Island, Lee County (n = 3), Volusia County (n = 1), St. Lucie County (n = 1), Indian River County (n = 1), Orange and/or Osceola Counties (n = 15). Collection of tortoises, except those from Orange and Osceola Counties, was opportunistic, and a result of submissions to the UF Wildlife Clinic, or other veterinary clinics. Some tortoises had exhibited signs of URTD, while others had been hit by automobiles. Tortoises from Orange and Osceola Counties were selected on the basis of clinical evaluations, ELISA, culture, and / or PCR results. Six clinically healthy animals were included in the latter group. Tortoises were evaluated for clinical signs of URTD (i.e., nasal and ocular discharge, palpebral edema, and conjunctivitis) and those exhibiting one or more signs of disease, with a past history of clinical signs, with positive culture or PCR results, or with a positive ELISA result, were designated as diseased. Tortoises were designated healthy if they were free of any history of or current signs of URTD and were culture, PCR, and ELISA negative. 30

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31 Figure 3-1. Locations in Florida from which gopher tortoises were obtained. 1 Alachua County, 2 Volusia County, 3 Orange/Osceola Counties, 4 Indian River County, 5 St. Lucie County, 6 Sanibel Island, Lee County. Necropsy and Histology Procedures Necropsy procedure and light microscopic evaluation of tissues were performed as detailed in Chapter 2. Gross necropsies were conducted on 21 tortoises. Multiple tissues were collected from 15 tortoises, heads and livers from six , and only heads from two. Microbial Isolation Flush and swab samples for Mycoplasma isolation were collected from the nasal passages and cavities of each tortoise and processed as described in Chapter 2. Swab specimens of the dorsal nasal cavities of 16 tortoises were collected for aerobic bacteria isolation and submitted to the Clinical Pathology Laboratory (CPL) of the College of Veterinary Medicine (CVM) at UF. Samples were cultured on Columbia blood agar and MacConkeys agar, and incubated at 37°C. Bacteria were identified utilizing the

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32 identification systems API 20E for enteric organisms and API NFT for non-enterics (BioMerieux Vitek, Hazelwood, MO, USA). Isolates of organisms consistent with Pasteurella were identified to species according to biochemical profiles listed for P. testudinis (Snipes and Biberstein 1982). Electron Microscopy Selected specimens were submitted to H. P. Adams of New Mexico State University, Las Cruces, for scanning and transmission electron microscopic preparation and evaluation. The left half of the bisected head of one healthy tortoise was prepared for ultrastructural evaluations. The nasal cavity was instilled with 2.5% glutaraldehyde in 0. 1 M phosphate buffer, then dissected out in its entirety, and selected areas were sampled. The tissues were dehydrated in an ascending series of ethanols and transferred to hexamethyldilisilazane for the final drying. The samples were sputter coated with gold and viewed by scanning electron microscopy (SEM). Samples from 1 1 diseased tortoises were collected for transmission electron microscopy (TEM). The nasal cavity tissue was removed from the underlying cartilaginous tissues and separated into anterior dorsal, anterior ventral, posterior dorsal and posterior ventral quadrants. Each quadrant was cut into 1 mm cubes and placed in 2.5% glutaraldehyde, and post-fixed in osmium tetroxide. Specimens were prepared for TEM by embedding in epon-araldite and sectioning with an ultramicrotome. Thick sections were stained with toluidine blue and examined by light microscopy. Ultrathin sections were placed on copper grids, stained with uranyl acetate and lead citrate, and examined with a Hitachi H7000 transmission electron microscope.

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33 Results Based on clinical evaluations and diagnostic tests, eight tortoises were classified as healthy and 15 as affected by URTD. Infected tortoises were from Orange / Osceola Counties (n = 10), Lee County (n = 3), Indian River County (« = 1) and St. Lucie County (n = 1). Histological findings for the two groups will be discussed separately. Normal Anatomy and Histology The external nares opened into ventro-lateral depression, and were continuous with large dorsal nasal cavities (Figure 3-2). Right and left dorsal nasal cavities were separated by a cartilaginous septum Each nasal cavity was bisected by a ridge, forming anterior and posterior compartments. Ventrally, the nasal passageways were continuous with the choanae (internal nares), which opened into the palatine region of the dorsal oral cavity. The integument continued through the external nares into a short vestibule, which was initially lined by keratinized stratified squamous epithelium That epithelium abruptly changed to mucous glandular epithelium, which lined the nasal passageway throughout its length. Interspersed among the mucous epithelial cells were ciliated epithelial cells. The ventro-lateral depression was lined primarily by mucous and ciliated epithelial cells (Figure 3-3a). Both anterior and posterior dorsal nasal chambers were lined by a multilayered olfactory epithelium with occasional mucous cells (Figure 3-3b). Numerous serous and mucous glands, vessels, nerve bundles, and clusters of melanophores were present in the connective tissue surrounding the nasal cavities. Small focal aggregates of lymphoid cells were seen in the submucosa.

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Figure 3-2. Diagrammatic representation of the interior of a gopher tortoise head sectioned longitudinally, illustrating the relationship of the nasal cavity to the external and internal nares. Approximately 3x life size. Drawing by L. Mallory.

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a Figure 3-3. Photomicrographs of normal gopher tortoise upper respiratory tract tissues. Hematoxylin and eosin staining, 320x. a) Photomicrograph of the ventrolateral depression demonstrating the ciliated mucous epithelium; b) Photomicrograph of the dorsal nasal cavity demonstrating the multilayered olfactory epithelium Photographs by E. R. Jacobson.

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36 The bilateral thymus glands were difficult to find in healthy tortoises and were located cranial to the base of the heart, at the branching of the subclavian and carotid arteries. Grossly, the thymus was multilobulated. Histologically, there was a typically dark staining cortex and a lighter staining medulla. The cortex contained densely packed thymocytes. In the medulla there were significantly fewer cells including thymocytes, as well as thymic epithelial cells, myoid cells, and heterophils. The spleen was located on the right side, between the proximal duodenum and transverse colon and was associated closely with pancreatic tissue. Histologically, spleens were composed of distinct areas of white and red pulp. White pulp consisted of collections of lymphoid tissue surrounding blood vessels. Red pulp, located between the perivascular collections of the white pulp, included red blood cells within sinusoids and small numbers of lymphocytes. The thyroid was located at the base of the heart. In one of the healthy tortoises, the thyroid was enlarged, with multifocal areas of follicular epithelial cell hyperplasia. In all tortoises, follicles varied in size, with many having numerous red blood cells in the colloid and either intraepithelial or supra-epithelial vacuoles. Multiple foreign bodies were seen in the submucosa of the glottis of two tortoises, in the tongue of one tortoise, and in the buccal salivary gland of one tortoise. The foreign bodies were consistent with plant material. Lymphoid aggregates were scattered throughout the esophagus, small intestine, large intestine and cloaca, and also were present in the connective tissue surrounding the mental (chin) glands. Of the healthy tortoises that were examined fully at necropsy, three were males and three were females. In one male, multifocal areas of mineralization of seminiferous tubules

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37 were seen. Golden brown granules were scattered throughout the kidney within renal tubular epithelial cells of all tortoises. Mononuclear cells containing similar appearing granules were within the renal interstitium and the interstitium of the testes. Pathologic Findings Gross examination of heads of diseased tortoises revealed rninimal to large amounts of exudate within the nasal cavity and nasal passageways. Histologically, of the 15 heads, one had no changes, two had mild changes, seven had moderate changes, and five had severe inflammatory changes. In the tortoises with minimal changes, mild mucosal hyperplasia and slightly increased lymphoid aggregates were seen in the nasal passage and ventro-lateral depression. In those tortoises with moderate changes, the olfactory epithelia were usually normal, with only focal or mild changes in the submucosa. Changes generally were confined to the nasal passage and the ventral aspects of the nasal cavities, and consisted of mucosal epithelial and lymphoid hyperplasia, with infiltration of mononuclear cells and heterophils. In some tortoises with moderate inflammation, basal cell proliferation and loss of cilia could be detected. In the tortoises with severe inflammatory changes, there were lymphoid aggregates around submucosal glands, with glandular epithelial hyperplasia. The normal mucosal architecture was replaced by infiltrates of mononuclear cells and heterophils. The olfactory mucosa was replaced with proliferating mucous epithelial cells (Figure 3-4). Proliferating basal cells projected into the underlying lamina propria of some tortoises. Exudate, consisting of sloughed epithelial cells and inflammatory cells, was found in the nasal cavity lumen. Infected tortoises had larger lymphoid aggregates in the submucosa of the glottis; several tortoises had focal areas of epithelial cell proliferation. Basal cell proliferation and

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38 Figure 3-4. Photomicrograph of the nasal cavity tissues of a gopher tortoise with upper respiratory tract disease. The changes were classified as severe, with aggregates of lymphoid cells in the submucosa, proliferation of the basal cells, and dysplasia of the mucosal epithelium. Hematoxylin and eosin staining, 320x. Photograph by E. R. Jacobson.

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39 submucosal lymphoid hyperplasia were seen in the glottis of one tortoise. The hyperplasia extended into the cranial tracheal epithelium. In that tortoise, there also were multifocal areas of epithelial cell hyperplasia in the lung. Five other diseased tortoises had focal to multifocal lymphoid aggregates in the lung interstitia. The gastrointestinal tracts tended to have increased numbers and larger lymphoid aggregates in the submucosa compared to those of clinically healthy tortoises. In one tortoise there was mucous cell hyperplasia of the colon and in another there was a severe colitis with basal epithelial cell hyperplasia and submucosal lymphoid hyperplasia, with infiltrates of large numbers of heterophils. Four other tortoises had increased lymphoid aggregates in the esophagus, stomach and/or small intestine. Three additional tortoises had increased lymphoid aggregates in the submucosa of the cloaca. The kidneys of all diseased tortoises contained golden brown pigment granules within renal epithelial cells. Hepatocytes of most tortoises contained similar granules, and pathologic changes were seen in the livers of nine tortoises. In seven, there were increased numbers and size of melanomacrophages in the liver and increased amounts of golden brown granules. One tortoise had cuffing of the central vein by lymphocytes and heterophils; another had aggregates of lymphocytes and melanomacrophages. Electron micrographs of tissues of one tortoise from Sanibel Island and the tortoise from Indian River County demonstrated organisms consistent with Mycoplasma on the surface of the nasal mucosa (Figure 3-5). Associated epithelial cells had vacuolated cytoplasm and inflammatory cell infiltrates were present in the mucosa. Increased numbers of mucous epithelial cells were seen, consistent with light microscopic findings.

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ELISA and PCR Results ELISA and PCR results for both healthy and diseased tortoises are presented in Table 3-1. All healthy gopher tortoises were seronegative for antibody specific against M. agassizii. Twelve diseased tortoises were seropositive, two were suspect, and one was seronegative. All healthy tortoises were PCR negative while 1 1 diseased tortoises were PCR-positive for Mycoplasma in nasal aspirates. Four clinically healthy tortoises that had negative culture and PCR results from nasal passage flush and swab samples had positive cultures and / or PCR results for samples from the nasal cavities. Microbial Isolation Results The results of Mycoplasma and aerobic microbial cultures of the upper respiratory tract (URT) are presented in Tables 31 and 3-2. Mycoplasma was not cultured from the URT of any healthy tortoise. Mycoplasma was cultured from the URT of 1 1 tortoises with URTD. The aerobic microbial isolates of healthy tortoises consisted primarily of members of the genera Staphylococcus, Streptococcus, and Coryne bacterium; a few Gram-negative rods were isolated. A greater number of Gram-negative species were isolated from the nasal cavities of tortoises with URTD, and those isolates made up a greater proportion of the isolates. Pasteurella testudinis was isolated from five diseased tortoises; in two it represented the major aerobic isolate. Discussion Gopher tortoises with clinical signs of URTD and evidence of exposure to Mycoplasma agassizii were obtained from multiple sites in Florida. The light microscopic

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Figure 3-5. Transmission electron photomicrograph of the nasal cavity mucosa of a gopher tortoise with upper respiratory tract disease. Organisms consistent with Mycoplasma (arrow) can be seen in close association with host cell membranes. Magnification 18,000x. Photograph by H. P. Adams.

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42 Table 3-1. Summary of ELISA, PCR, culture, and nasal histopathology results from necropsied gopher tortoises from various locations in Florida, for determining infection with Mycoplasma agassizii. Group Clinical Signs ELISA PCR Mycoplasma Culture Histopathology Normal (« = 8) 0% 0% 0% 0% 0% Natural Infection (« =15) 60% 85% 92% 80% 92% Table 3-2. Summary of aerobic culture results from nasal cavity swabs of gopher tortoises from Florida. Percent Group Growth Species of isolates Normal very scant to Staphylococcus spp. 45 (n = 5) moderate Gram-negative rods 25 a-hemolytic Streptococcus sp. 10 non-hemolytic Streptococcus spp. 10 Corynebacterium spp. 10 Natural very scant to Coryne bacterium spp. 33 Infection heavy Pasteurella testudinis 18 (n=ll) Staphylococcus spp. 16 Gram-negative rods 15 Micrococcus sp. 11 Flavobacterium, Pseudomonas, 7 Lactobacillus, and others

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43 changes in the upper respiratory tract of the gopher tortoises were similar to inflammatory and dysplastic changes reported for desert tortoises with URTD (Jacobson et al. 1991, Brown et al. 1994, Jacobson et al. 1995). However, inflammationa and epithelial proliferation around the glottis, tracheitis, and proliferative pneumonia have not been seen in desert tortoises. Additionally, two diseased tortoises had proliferation of the colonic mucosal epithelium Those changes have not been seen in desert tortoises with URTD. By electron microscopy, organisms consistent with Mycoplasma were demonstrated on the nasal mucosal surfaces of two tortoises. Other than bacteria, no infectious agents were demonstrated in or on nasal cavity mucosa by electron microscopy. Eleven diseased gopher tortoises were PCR positive and M. agassizii was cultured from the upper respiratory tract of 1 1 diseased tortoises examined. These results support the hypothesis that M. agassizii is a cause of URTD in gopher tortoises in Florida. The greater number of species and increased proportion of Gram-negative bacteria isolated from diseased tortoises could indicate that conditions in the upper respiratory tract of diseased tortoises are more favorable for the growth of those species, or that tortoises infected with M. agassizii are more susceptibile to opportunistic pathogens. The positive culture and / or PCR results from nasal cavity samples (obtained at necropsy) of four clinically healthy tortoises with negative nasal passage flushes and swabs support the hypothesis that tortoises can harbor the organism without showing clinical signs or shedding bacteria. Such animals may recrudesce under stressful conditions, begin shedding bacteria, and become infective to other tortoises. Nine gopher tortoises exhibited pathologic changes in the livers, although the significance of these changes is not understood currently. The mycoplasma may release

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toxic compounds or induce the production of compounds by the tortoises that cause damage to the liver. Desert tortoises with URTD also show changes in liver tissue (B. L. Homer, pers. comm.), and some exhibit altered nitrogen metabolism (B. Henen, pers. comm.). The altered nitrogen metabolism may be due to behavioral changes leading to decreased foraging rates or efficiency, or it may be due to direct effects of the Mycoplasma infection, but the mechanism is not yet understood. Although only one of nine experimentally infected captive gopher tortoises significantly decreased its intake of vegetables (G. S. McLaughlin, unpub. data), and that decrease was temporary, wild tortoises may alter their behavior patterns to a greater degree. Alternatively, secondary infections by other bacteria may be the proximate cause of liver damage. However, no evidence of primary or secondary bacterial infection (i.e., necrosis) was seen histologically. By the ELISA, none of eight clinically healthy tortoises and 11 of 1 5 diseased tortoises had antibodies against M agassizii, indicating previous exposure. Tortoises with negative ELISA results may have been in the early stages of the disease, when increased antibody levels had not occurred or were not detectable. The four diseased, seronegative, tortoises may have been infected with another agent. Westhouse et al. (1996) implicated an iridovirus as the cause of pneumonia, tracheitis, pharyngitis, and esophagitis in a gopher tortoise from Sanibel Island. The virus was readily detectable on both light and electron microscopy. It is not known if attempts were made to culture mycoplasma, or if an ELISA was run on a plasma sample. No indications of viral infections were seen in the tortoises examined for the current study. Investigators recently have found seropositive gopher tortoises in Georgia (B. Raphael pers. comm.), seropositive, clinically ill, and/or PCR positive tortoises at a site in

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45 northeast Florida, and seropositive tortoises at a site in Mississippi (D. M. Epperson, pers. comm.). Although many Florida and Georgia tortoise populations are fairly large, those in Mississippi are more restricted and are on the federal endangered species list. Outbreaks of URTD in populations with limited recruitment and no nearby source populations could contribute to severe declines in numbers and, possibly, local extinctions. Further studies, on a range-wide basis, need to be conducted to determine the distribution and potential impacts of URTD on gopher tortoise populations. In addition, further research is necessary regarding the pathogenesis of URTD. While findings of liver, kidney, and intestinal tract lesions are interesting, they do not elucidate the mechanisms by which the changes are caused. Changes have been found in hormonal profiles of some infected desert tortoises (Rostal et al. 1996), which could lead to altered foraging and reproductive behavior, as well as decreased reproductive potential. If foraging behavior is affected, food and water intake might be reduced, which could affect liver and kidney functions. Both direct and indirect pathogenic mechanisms need to be studied in order to better predict the effects of URTD on tortoises.

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CHAPTER 4 EFFECTS OF REPEATED EXPOSURE ON SEROPOSITIVE ADULTS Introduction In order to properly evaluate the results of the diagnostic tests and incorporate those findings into management and conservation plans, epidemiological questions, including the response of seropositive, asymptomatic tortoises to subsequent exposure to the agent, must be addressed. Although vaccines have been developed for some mycoplasmal diseases (Ellison et al. 1992, Lai et al. 1996, Markham et al. 1996), they are usually not fully protective (e.g., Djordjevic et al. 1996, Kleven et al. 1996, Mohan et al. 1996, Washburn and Weaver 1996), and many natural infections by mycoplasmas do not engender a protective host immune response. The immune response is actually essential to the development of lesions, and infected animals are susceptible to repeated infection (Simecka et al. 1992). Due to the immunopathology, the disease may be more severe on subsequent exposure than on initial infection. I designed an experiment to test the hypothesis that tortoises that have produced antibodies against M agassizii are protected against reinfection with the organism and subsequent development of URTD. 46

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47 Methods Acquisition of tortoises, intake and husbandry procedures, clinical assessments, sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed in Chapter 2. Statistical Analyses The onset and severity of clinical signs of URTD, and ELISA, PCR, and culture results for the challenge tortoises were compared to those for control and naive tortoises using the SAS system (SAS Institute, 1988). Data from an additional naive animal infection experiment were included after it was determined that the data did not differ significantly from those collected from naive animals in this study. This ensured large enough sample sizes for meaningful comparisons at more time points. Differences in the severity of clinical signs, histologic lesions, and ELISA data among the three treatment groups were compared by an analyses of variance-type logistic regression using maximum likelihood estimators to compensate statistically for the different number of tortoises in each group at each sampling date. Percentages of tortoises showing clinical signs at different time points were compared by Fisher's Exact test, with a P value of 0.05 accepted as significant. Experimental Design Four groups of tortoises were established. Three groups [control (n = 6), naive (n = 11), and sentinel (n = 2)] had no history of exposure to M. agassizii, while the fourth, or challenge group (n = 8), had previous history of exposure as indicated by a positive result

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48 on the ELISA. Initially, no tortoise in any group had clinical signs of URTD, or positive culture or PCR results. One tortoise originally slated for inclusion in the challenge group (i.e., seropositive, but clinically, culture, and PCR negative upon arrival) developed clinical signs before inoculation and was eliminated from the study. Approximately 1 mo following arrival, the controls were sham inoculated intranasally with 100 ul sterile SP4 broth in each naris, and tortoises in the naive and challenge groups were inoculated in each naris with 100 ul of SP4 broth containing approximately 10 4 colony forming units (CFU) ofM agassizii strain 723, for a total dose of 10 8 CFU. The 723 isolate was obtained originally from a clinically ill tortoise from Sanibel Island, Lee County, Florida. The sentinel tortoises were captured, but received no other treatment. Following inoculation, observations of all tortoises were attempted daily to determine the onset and sequence of clinical signs. Behavior also was monitored. At 2 4 wk intervals post-inoculation (PI), tortoises were trapped, examined, and weighed, then tranquilized and blood, nasal flush and nasal swab samples were collected. Samples were processed as previously described. A total of 22 tortoises was examined at necropsy and histologically. Four control and nine naive tortoises were euthanatised and necropsied in October, 1994, before undergoing winter dormancy; seven challenge and two naive animals were euthanatised and necropsied in March, 1995, after emergence. Complete necropsies (n = 15) were performed on two control, six naive and seven challenge tortoises; only heads were examined on the other animals (n = 7).

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49 Results Clinical Signs Of the six control group tortoises, none showed consistent clinical signs nor had positive culture, PCR, or ELISA results. At each of three time points, one control tortoise exhibited mild ocular signs that were probably associated with environmental conditions. One of two sentinels developed clinical signs at 12 wk PI, although she had not been inoculated with culture or medium. Of the naive tortoises, 67% exhibited clinical signs beginning 2-3 wk PI, 79% were clinically ill by 8 wk PI, and 94% had shown signs by 16 wk PI (Table 4-1). One naive tortoise never exhibited clinical signs. All seven challenged tortoises developed clinical signs of URTD before dormancy. Five (71%) challenged tortoises and one naive tortoise exhibited clinical signs soon after emergence. No tortoises became moribund or died during the study. At 2 wk PI, six challenge and 1 7 naive tortoises were examined. Challenged tortoises had higher total clinical sign scores than naive tortoises (2.6 vs. 0.5, P < 0.001), and higher scores for nasal discharge ( 1.5 vs. 0. 1, P < 0.001), ocular discharge (0.8 vs. 0, P < 0.02), and palpebral edema (1.5 vs. 0.6, P < 0.02) (Table 4-1). Significantly more challenge than naive tortoises exhibited nasal and ocular discharge (67 vs. 6%, P < 0.01; 50 vs. 0%, P < 0.02; respectively) (Figure 4-1). No consistent differences were seen between the two groups at later sampling times (Figure 4-2).

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50 Table 4-1. Percentages of naive and challenge tortoises positive for each clinical sign of URTD, and mean clinical sign scores. Scores < 1.5 are classified as slight, >1.5 and <2.5 as moderate, and >2.5 as severe. ND = nasal discharge, OD = ocular discharge, ED = palpebral edema, CJ = conjunctivitis. P < 0.05 indicates significant difference between groups. Time % Positive Mean Score Sien Naive Challenge p Naive Ch alienee p Z WK Total o / 8"5 u.ozo o 5 U. J Z.O o ooo u.uuu \rn a o a 7 o / 0 007 u.uu / 0 1 U. 1 1 5 1. J O 000 cxn A U so 0 011 U.U 1 1 A u 0 8 U.O U.U1 j pn 5Q jy 8"? OJ O 7A0 O A U.O 1 5 O 0 1 9 U.U1Z 57 O 740 U. J4U O A U.O 1 o l.U O 745 U. j4 J 4 wk Total 1 Old 1 70 ly oU 1 000 1 .uuu 1 5 1 Q l.y O 495 U.4Z J \m JU nO 1 000 0 8 U. o 1 9 1 .z O 9 54 U.Z nn HO zu U. JOO 0 8 U. o 0 fx U.O U.OOO cn 11 u 59 tu 1 000 1 .uuu 0 7 u. / 0 8 U. O 0 8o4 U.oOH CJ SO AO OU 1 000 1 .uuu 1 0 1 .u 0 8 U. o 0 o70 U.O /U 6 wk Total i oxai 1 oo 1UU 1 ooo 1 .uuu 7 1 Z. 1 9 7 Z. / O 990 u.zzv n7 O / OJ u.ozo 1 1 1 . 1 1 Q O 040 U.UHU on VJIJ 44 j / 0 f\TX U.O / J 0 5 U. D 1 0 l.U 0 118 U. 118 en 7? 7 1 1 000 1 .uuu 1 0 1 .u 0 Q U. y 0 o59 u.o jz 80 O" 4^ HJ U.UJZ 1 4 0 7 u. / 0 055 U.U J J o WK i otai 89 oZ 1 ooo 1 .uuu 7 A Z.O Z.4 A ^OO U.OOO Kin JN1J 71 / 1 71 1 ooo l.UUU 1 A 1.4 i a 1.0 A £77 U.O / / an 5/1 57 J / l.UUU A A u.y a n U. / A A HC\ en 71 7 1 l.UUU 1. 1 i ^ l.i U.Oj4 71 90 zy O 075 U.U / J 1 C 1 . J A A U.O A AA/1 U.UU4 19 wk Total 83 83 1.000 2.4 2.4 0.989 ND 62 71 0.642 1.5 1.6 0.836 OD 54 57 0.676 1.0 0.7 0.418 ED 67 71 0.667 0.7 1.3 0.068 CJ 79 29 0.198 1.1 0.6 0.105 Nx Total 84 83 1.000 2.6 2.4 0.744 ND 79 71 0.646 1.4 1.6 0.733 OD 79 57 0.340 1.2 0.7 0.123 ED 58 71 0.668 1.0 1.3 0.283 CJ 63 29 0.190 1.3 0.6 0.036

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i = -= Q O H 09 o o o o 00 o o o o o m o o uoxpajrajsod s^sa^ 2 'saijisoj sssiojjox %

PAGE 61

52 to

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53 Culture and PCR Results The sentinel tortoise that became ill also had positive culture and PCR results. All inoculated tortoises had at least two positive culture results, and 17 of 18 had positive cultures from samples collected at necropsy. Mycoplasmal DNA was detected by PCR analysis from each inoculated tortoise, including those that did not exhibit clinical signs of URTD when sampled. By RFLP analysis, all isolates corresponded to M. agassizii. ELISA Results There was no anti-M agassizii antibody response by the control tortoises to sham inoculation, although one sentinel had seroconverted by 4 wk PI. All inoculated tortoises seroconverted or had significantly increased ELISA values. ELISA values were greater for challenge than naive tortoises at each time point (Table 4-2, Figure 4-3). A significant increase in ELISA values of challenged tortoises was observed by 4 wk PI (mean ratio of samples to negative control of 6.3 vs. 3.3 at 0 wk, P < 0.05), and seroconversion of naive tortoises was observed by 6 wk PI (ratio of 3.9 vs. 1.5 at 0 wk, P < 0.05). The ratio of the challenge to the naive ELISAs increased at 2 and 4 wk PI, then declined. Table 4-2. Least-squares mean ELISA values for naive and challenge tortoises. P < 0.05 indicates a significant difference between groups. Time Naive Challenge P 0 weeks pi 0.2049 0.4707 0.0437 2 weeks pi 0.2235 0.5925 0.0126 4 weeks pi 0.2439 0.8848 0.0001 6 weeks pi 0.5464 1.5254 0.0001 8 weeks pi 0.8314 1.8219 0.0001 12 weeks pi 1.4062 1.9839 0.0001 Necropsy 1.6796 2.2889 0.0001

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54 00 o ,4> O O o o o © o o ura got 7 © souBqjosqy "5 "5 ^ 1/1 J -S tt £>
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55 Histology Results The upper respiratory tracts of the four control tortoises examined at necropsy had normal histologic appearances. In contrast, all tortoises inoculated with mycoplasma showed lesions similar to those seen in naturally occurring URTD. All challenge tortoises had moderate to severe inflammation and changes in the epithelium and submucosa. Three naive tortoises had minimal lesions and eight had moderate to severe abnormalities.. Lesions were consistently seen in the ventrolateral depression of the nasal cavity, a region immediately caudal to the vestibule (see Chapter 3). Discussion No tortoises inoculated with sterile medium developed clinical signs or histopathologic lesions, indicating that lesions were not a result of the mechanical effect of the medium on the tissues, nor of host inflammatory response to the medium. Experimentally infected naive animals did not seroconvert until 6 to 8 weeks PI. Because the sentinel that became ill and seroconverted was sampled only on arrival and 8 wk following arrival (4 wk PI for the inoculated tortoises), the timing of its seroconversion is uncertain. No known clinically ill tortoises were transported or held with the experimental animals once they left the Orange County holding facility, and the sentinel was not held in the facilities or with other tortoises during the time tortoises were being inoculated. Additionally, in an experiment involving the transfer of healthy tortoises into pens immediately following the removal of ill tortoises, there was no indication of environmental transfer, although the sample sizes were very small (see Chapter 7).

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56 Therefore, the sentinel whose seroconversion was detected 8 wk after arrival probably was exposed to M. agassizii just before transport to UF. Tortoises harboring M agassizii may not show clinical signs, may exhibit mild signs, or may show signs only intermittently. Because tortoises not showing any ocular signs or nasal discharge sometimes have positive culture and PCR results, the organism may be transmissible from asymptomatic tortoises under appropriate conditions. Due to unavoidable constraints on sampling live animals, I cannot exclude the possibility that clinically healthy, seropositive, culture and PCR negative animals harbor the bacteria. As shown by the two tortoises (the sentinel and the tortoise that was eliminated from the study) that initially had negative culture and PCR results but developed disease without being inoculated experimentally, some infections may go through an extended latent period with low numbers of organisms in the nasal passages, or animals that appear to have cleared the organism may, in fact, have not. The clinical response of challenged animals was more rapid and severe than that of naive animals, indicating that no protection was conferred by previous exposure to the organism This is consistent with some other mycoplasmal diseases in which the immune response confers limited or no protection (Ellison et al. 1992), or contributes to pathogenesis, such as arthritis in fowl caused by M. synoviae (Kume et al. 1977), conjunctivitis in cattle caused by M. bovoculi (Rosenbusch 1987), and pneumonia in humans caused by M pneumoniae (Krause and Taylor-Robinson 1992). Tully et al. (1995) have found that some mycoplasmal surface proteins share sequence and structural homologies with vertebrate proteins, and suggest that these may play a role in eliciting autoimmune responses. Repeated exposure to mycoplasmal proteins that resemble a

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57 host's proteins may sensitize the host and induce an autoimmune response, leading to chronic manifestations of disease even if the primary etiologic agent is cleared. Although actively produced antibodies apparently do not confer protection to adult tortoises, it is unknown if passively transferred antibodies (see Chapter 6) provide protection against URTD for hatchlings. Preliminary observations indicate that seronegative hatchlings are at least as susceptible to infection as adults, and that the disease progresses more rapidly, with high morbidity in the first 6 mo PI (see Chapter 5). Desert tortoise hatchlings exposed to infected adults developed clinical signs of URTD and suffered extremely high mortality rates (5 1 of 69, or 74%, of those with clinical signs died) (Oftedal et al. 1996). Mycoplasma agassizii was isolated from necropsied individuals, and severe inflammation was seen histologically. The immune status of the hatchlings and parents was unknown prior to the development of the outbreak, so no inferences can be drawn regarding the potential role of maternal antibodies in the course of URTD in hatchlings. Because cellular components probably play a major role in the pathogenesis of URTD and those are not transferred vertically in reptiles, the maternal antibodies may provide some protection. It would be interesting to explore the question of increased susceptibility to or severity of disease vs. protection from disease in hatchlings that have received maternal antibodies against M. agassizii. The immune response stimulated by previous exposure to M. agassizii did not prevent reinfection or ameliorate signs of URTD after subsequent inoculation. The onset of clinical signs was, in fact, more rapid and the initial severity was greater than in first infections. Development of specific antibodies against M. agassizii did not ensure clearance of the organism by the tortoises, either on initial or subsequent exposure. Based

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58 on this study, tortoises testing positive for antibodies against M agassizii cannot be considered good candidates for release in repatriation, restocking, or relocation efforts. It may be acceptable to release seropositive tortoises in areas that already have seropositive animlas, as long as the overall prevalence is not increased significantly. Seropositive tortoises may provide desirable genetic material (see Chapter 6), and would be valuable as research subjects to study interactions with other disease agents and the effects of repeated exposure on metabolism, autoimmune responses, reproduction, and survival.

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CHAPTER 5 HORIZONTAL TRANSMISSION OF MYCOPLASMA AGASSIZII Introduction In order to develop management and conservation plans that incorporate the potential of disease to affect populations, epidemiologic questions must be addressed. Of particular concern are the probability of transmission of the disease organism from one tortoise to another (horizontal transmission), the rate of spread within populations, and the potential for spread to nearby populations. The probability of transmission may be related to a tortoise's clinical and culture or PCR status. Infected tortoises may be culture and/or PCR positive without showing clinical signs. Conversely, it is sometimes difficult to detect bacteria in clinically ill tortoises. I designed an experiment to test the hypothesis that horizontal transmission of M. agassizii will occur only from those individuals that are clinically ill, and PCR and/or culture positive. Methods Acquisition of tortoises, intake and husbandry procedures, clinical assessments, sampling methods, ELISA and PCR procedures, and necropsy procedure were as detailed in Chapter 2. 59

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60 Experimental Design Fifteen pairs consisting of one male and one female were established in August, 1994. Due to the limited availability of seropositive and/or clinically ill tortoises, the sample design was not balanced. Five pairs of asymptomatic, ELISA-, PCR-, and culturenegative control tortoises were established as controls. The other ten pairs consisted of a clinically healthy, ELISA-, PCR-, and culture-negative tortoise that had been at UF since April, 1994 (resident), and an ELISA-positive or -suspect (irrespective of PCR or culture status) tortoise of the opposite sex and similar size (Figure 5-1, Table 51). One resident female was paired with an ELISA-negative, but clinically ill, cultureand PCR-positive male. The serosuspect and two seropositive tortoises were clinically healthy, and had negative culture and PCR results. The rermining six seropositive tortoises showed moderate clinical signs of illness. Three of those six were cultureand PCR-negative, one was cultureand PCR-positive, one was culture-positive but PCR-negative, and one was culture-negative and PCR-positive. Behavioral observations and clinical signs of URTD were recorded opportunistically (see Chapter 2), and blood and nasal flush samples were collected from all tortoises in August and October, 1994, and in March and August, 1995. In 1996, samples were collected in February or March, and in July or August. Clinical signs and weights were recorded, and photographs were taken at each sampling time. Data were analysed using Chi-square and Fisher Exact tests for differences in proportions of tortoises becoming infected under different exposure conditions. Infected animals were euthanatized in July, 1996, except as noted below, and uninfected tortoises were released pursuant to Florida Game and Fresh Water Fish Commission permit number WR96224.

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61 Seronegative Clinically 111 Culture & PCR Positive n= I Seropositive or Suspect n = 9 Clinically Healthy Culture & PCR Negative n = 3 Clinically 111 // = 6 Culture & PCR Negative 3 Culture & PCR Positive 1 Culture or PCR Positive n = 2 Figure 5-1. Flow chart showing initial distribution of seropositive, clinically ill, culture and PCR positive tortoises.

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Results General Observations Observations indicated varying degrees of interaction between pair members. Several males were excluded from the original burrow in the pen by their female partners. Until one of the two dug a new burrow, those males, particularly 125, 185, 201, and 235, spent most of the day in a corner of the pen, under the shade cloth, and sometimes spent the night out of the burrow. Other pairs spent considerable time in courtship, or foraging in close proximity to one another. In March, 1995, five hatchling tortoises were found in pen Dl. The female (135) may have been gravid or storing sperm when she arrived at the UF facility in April 1994, and laid the clutch during the summer of 1994. Alternatively, she may have copulated with the male (241) and ovulated shortly after the pairing in August 1994. Based on gopher tortoise physiology (Taylor 1982), however, I believe the former explanation is more likely. One hatchling was removed and euthanatised due to congenital abnormalities; the other four were left in the pen. Three tortoises were euthanatised in June 1995 (Table 5-1). One, number 287 (pen D10), was an initially seropositive, clinically ill, culture and PCR negative female whose partner (125) had not become ill. She failed to reproduce, and a large mass could be seen on radiographs. At necropsy, a large urolith (bladder stone), approximately 5 cm in diameter, was removed. The second, number 213 (pen D5), was an initially seropositive, clinically ill, culture and PCR positive female whose partner had become ill within one month of pairing. She failed to lay six eggs, and on exploratory surgery it was discovered that both oviducts had ruptured and the eggs were free in the coelomic cavity.

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65 She was euthanatised without recovery from anesthesia. Her partner, number 185, was clinically ill, lethargic, and losing weight, and was euthanatised shortly thereafter. One healthy, seronegative male tortoise (144, pen D2), paired with a seropositive female (275) disappeared after March 1995 (Table 5-1). He may have been overturned during interactions with the female and been pushed into the burrow, where he would have been unable to right himself. At the time of his disappearance, he had exhibited no indications that transmission of M. agassizii had occurred. The male tortoise from pen D6 (183) also disappeared, and may have met a fate similar to that of 144. His disappearance may have been related to agonistic interactions with any of the tortoises originally in pens Dl, D6, D7, or D8, as discussed below. Due to the nature of tortoise burrows, it was impossible to determine unequivocally the fates of the animals that disappeared. Approximately 10-1 1 months after pairing, in June and July 1995, the burrowing activities of the large number of tortoises in the small area resulted in the communication of several pens (Figure 5-2). Tortoises in pens Dl, D6, D7, and D8 moved regularly between pens and had numerous agonistic encounters with one another. The four tortoises in pens D3 and D4 also moved between pens and had frequent interactions. Although tortoises from pens D9 and D10 also interacted with each other, aggressive interactions were not observed. One control male (147, pen A2), burrowed into an adjacent pen (A7) containing an experimentally infected ill tortoise, and moved between the two pens over several months. Neither the control female (577) nor the third tortoise was observed to switch pens. The study was originally designed to last for 12-14 mo, but when the animals began moving between pens, it was extended to 24 mo in order to collect more data.

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66 Figure 5-2. Burrow map showing interconnections among pens Dl, D6, D7, and D8; between D3 and D4, and between D9 and D10. • Burrow entrances. Evidence of Transmission of Mycoplasma agassizii The male control tortoise (147) became ill, seroconverted, and had positive culture and PCR results (Table 5-1). His partner became clinically ill and seroconverted after he returned to his original pen. No other control tortoises became ill or seroconverted. The D-section burrowing activities mentioned above resulted in the exposure of all tortoises in pens Dl and D7, which until that time had been clinically healthy, to clinically ill tortoises with positive culture and/or PCR results. Three became clinically ill, and had increased ELISA readings, with one of two initially negative tortoises seroconverting. Those activities also resulted in the exposure of the tortoises in pen D8 to clinically ill, culture and PCR positive tortoises. The seronegative male (130) became ill, had positive culture results, and seroconverted; the clinically ill, seropositive female (257), which had been culture and PCR negative until that time, also became culture positive. All four hatchlings in pen Dl became severely ill, seroconverted, and had positive culture results. All were euthanatised, and pure cultures of M. agassizii were recovered from the nasal cavities and conjunctiva on necropsy.

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67 The initially seronegative, clinically ill, culture and PCR positive male (183) had seroconverted by October 1994. Based on his original samples from July 1994, and the ELISA values of his October samples, he probably seroconverted within 2 wk of pairing, and was considered as ELISA positive for the analyses (see below). In the ten experimental pairs, eight initially asymptomatic members, two of which were originally ELISA-positive and six of which were originally ELISA-negative, became moderately ill with signs of URTD and had increased ELISA values, with the latter six seroconverting. Two other tortoises (125 and 190) showed mild clinical signs that may have been associated with mechanical irritation or environmental conditions, but did not seroconvert. Of the four initially uninfected tortoises that showed no increase in antibody levels, three did not have positive culture or PCR results. One (190) had only mild ocular signs, and two (110 and 144) never showed clinical signs. The fourth (125), which consistently showed mild signs and had a suspect PCR result in October 1994, was exposed to a tortoise (287) that had positive samples only at necropsy. Based on observations, they had only limited interactions. The serosuspect female (205), who was in contact with both 1 10 and 125, showed occasional, transient, mild clinical signs, but never had positive cultures or PCR results, nor did her antibody levels increase. In fact at all sample times after August 1994, she was classified as seronegative. Of the 16 originally culture and PCR negative tortoises, 10 later had positive or suspect culture or PCR results. Eight were exposed to clinically ill, culture or PCR positive tortoises, seroconverted and had positive results, the ninth had positive culture results only from necropsy samples, and the last was the individual (125) discussed above. No tortoise became ill unless exposed to another ill tortoise.

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Transmission Probabilities Analysis of the data was complicated by the changing exposure status of the tortoises due to the housing problems discussed above. Therefore, analyses were carried out on three data sets. The first consisted of data from the first 10 mo (August 1994 June 1995) of the study, the second from the last 14 mo (June 1995 July 1996), and the third of the cumulative data. The first data set (Table 5-2) consisted of the original 15 pairs. The second set (Table 5-3) did not include those pairs in which transmission had already occurred, but only the control pairs and those whose exposure status had changed, leaving 1 1 observations. The third set (Table 5-4), with 22 observations, included some tortoises twice, as the observations of interest were the exposure events themselves, and not the pairs or individual tortoises. Although sample sizes were very small, and some cells had no observations, chi-squared tests of differences in proportions were run on the three data sets. The final set was collapsed on the three predictor variables (ELISA, clinical, and culture/PCR status; Table 5-5) to determine which had the most influence on transmission probability. Four exposure status classes were established based on ELISA classification, clinical signs, and culture or PCR status. Class assignment was based on a tortoise's status at any point during the time period of interest. Therefore, a tortoise's status could change between the first and last parts of the study. Each criteria was recorded as positive or negative, and transmission was recorded as yes or no. Transmission was classified very conservatively, with any tortoise exhibiting even mild, transient, clinical signs not obviously related to mechanical irritation (e.g., plant material in the eye or nose) or environmental conditions (e.g., being held overnight in a box in which the tortoise had

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69 urinated), or having one suspect culture or PCR result recorded as having been infected by horizontal transmission. The four classes were 1) ELISA, clinically, and culture/PCR negative (-/-/-), 2) ELISA positive, clinically and culture/PCR negative (+/-/-), 3) ELISA and clinically positive, and culture/PCR negative (+/+/-), and 4) ELISA, clinically, and culture/PCR positive (+/+/+). The first data set (Table 5-2) consisted of the following observations (class, yes, no): 1,0,5; 2,0,2; 3,2,2; and 4,4,0. The second set (Table 5-3), for the last 14 mo, was comprised of the following observations: 1,0,4; 2,0, 1; 3,0,1; and 4,5,0. The last, cumulative, data set, included the following observations: 1,0,5; 2,0,3; 3,2,3; and 4,9,0. Chi-square tests of association of exposure status and transmission were significant for all data sets, with 3 degrees of freedom (df) and P < 0.001 for all sets (Tables 5-2, 5-3, 5-4). Table 5-2. Contingency table for exposure status and transmission occurrence for the first 10 mo (August 1994 June 1995) of the gopher tortoise URTD pairing study. Exposure Status ELISA/Clinical/Culture or PCR Transmission Yes No -/-/0 5 +1-10 2 +/+/2 2 +/+/+ 4 0 X 2 , 3 df = 10.8, P 0.013, power = 0.80 Table 5-3. Contingency table for exposure status and transmission occurrence for the last 14 mo (June 1995 July 1996) of the gopher tortoise URTD pairing study. Exposure Status ELISA/Clinical/Culture or PCR Transmission Yes No -/-/0 4 +1-10 1 +1+10 1 +1+1+ 5 0 X 2 , 3 df = 1 1.0, P = 0.012, power = 0.81

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70 Table 5-4. Contingency table for exposure status and transmission occurrence for the duration (August 1994 June 1995) of the gopher tortoise URTD pairing study. Exposure Status ELISA/Clinical/Culture or PCR Transmission Yes No -/-/0 5 +1-10 3 +/+/2 3 +1+1+ 9 0 X 2 , 3 df = 17.2, P = 0.0006, power = 0.96 Table 5-5. Transmission probabilities and tests of significant differences of proportions for different exposure status classes of tortoises relative to upper respiratory tract disease. Exposure Status Transmission Yes No Probability of Transmission Fisher Exact Test, P value ELISA negative positive 0 5 11 6 0 0.65 0.018 Clinical illness negative positive 0 8 11 3 0 0.79 0.001 Culture / PCR negative positive 2 11 9 0 0.15 1.00 0.0002 Because transmission was defined very conservatively for the purposes of this study, two tortoises were categorized as having been infected via horizontal transmission based solely on the appearance of mild clinical signs. Neither animal seroconverted nor had positive or suspect culture or PCR results during the second half of the study. Both tortoises were housed with clinically ill, seropositive tortoises that were culture and PCR negative. If those tortoises were designated negative relative to transmission, the final

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71 data set was comprised of the following observations: 1,0,5; 2,0,3; 3,0,5; and 4,9,0. The chi-squared value was 22.0, with 3 df and P < 0.0001. When Table 5-4 was collapsed on ELISA status (Table 5-5), none of the tortoises in five ELISA negative pairs became ill or seroconverted, but of the 17 interactions involving ELISA positive tortoises, 1 1 resulted in transmission, for a probability of 0.65. Fisher Exact test of a difference in proportions was significant at P = 0.018. When collapsed on clinical status, none of eight interactions involving negative tortoises resulted in transmission, as defined above, while 11 of 14 (probability 0.79) involving clinically ill tortoises did (Fisher exact test, P = 0.001). If transmission was defined more liberally, and designated as positive only if a tortoise seroconverted and had clearly positive culture or PCR results, then nine of 14 (probability 0.64) interactions with clinically ill tortoises resulted in transmission (Fisher exact test, P < 0.006). When transmission was defined conservatively, for culture and/or PCR status, 2 of 13 culture/PCR negative class interactions resulted in transmission (probability 0. 15), while all of nine positive class pairings effected transmission (Fisher exact test, P 0.0002). If transmission was defined more liberally, then none of 13 culture/PCR negative interactions resulted in transmission, and the Fisher exact test was significant with P = 2\ 10" 6 . Discussion The unplanned movements of the tortoises resulted in more exposure events than initially planned, and possibly compromised some of the original experimental design. However, at least one pair remained in each exposure category and was followed for 24

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72 mo, twice as long as originally planned. Therefore, the intent of the study was not compromised by the tortoises' natural behavior. Although manifestation of latent infections could not be excluded absolutely, the results supported the hypothesis that M. agassizii was horizontally transmitted between adult tortoises, and between adult and hatchling tortoises. The route of transmission may have been direct contact, aerosol, or fomite transmission. Direct contact was the most likely route. If aerosol transmission occurred, it was probably over short distances, such as would be likely to lead to direct contact. Sentinel and control animals housed in pen groups that also housed ill tortoises did not become ill or seroconvert, indicating that aerosolized bacteria were unlikely to travel even relatively short differences or over low (0.5 m) barriers. Similarly, fomite transmission is unlikely to play a major role (see Chapter 7). However, because many tortoise interactions occur in burrows, it is difficult to assess the importance of the various modes. Although sample sizes were small, results supported the hypothesis that transmission is more likely when the "donor" is symptomatic, although tortoises without clinical signs may be infected and able to transmit the pathogen under appropriate conditions. After rainstorms, it is not uncommon to find two or more tortoises drinking from the same puddle (R. E. Ashton, pers. comm.; G.S. McLaughlin, pers. obs.). When tortoises drink, they often get water in their noses, which they then blow or "sneeze" out. An asymptomatic tortoise harboring bacteria may shed enough in this manner to infect a nearby conspecific via the aerosol route, or to leave an infective dose in the water or on nearby plants.

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73 A concurrent study (Brown et al. 1996b), in which gopher tortoises were experimentally infected with varying doses of M. agassizii, demonstrated that the infective dose of the 723 strain is very low, less than 10 CFU. However, an additional isolate failed to induce disease or elicit an antibody response. In previous research involving desert tortoises, Brown et al. (1994) demonstrated that exudate from ill tortoises elicited a stronger immune response and induced more severe disease in inoculated tortoises than pure cultures of M. agassizii. In addition to the mycoplasma, the exudate probably contains other species of bacteria (e.g., Pasteurella testudinis, Coryne bacterium sp.), as well as cellular components and cytokines that may elicit an immune response and contribute to the disease process. The two tortoises that exhibited mild signs without seroconverting or having positive culture and PCR results may have been responding to cellular or chemical components in the exudate from their partners, to environmetnal conditions, or to other species or strains of bacteria that were only mildly pathogenic. Because conjunctival swabs were not collected from these animals, and aerobic cultures were not performed, those possibilities were not addressed in this study. Further research regarding transmission probabilities and modes is needed. Larger sample sizes and balanced designs would allow more rigorous statistical analyses. However, establishing and maintaining such designs would be difficult, as tortoises' clinical, culture, and PCR status can vary over both the short and long term (e.g., see Chapter 4). Experiments entailing differences in pen design could be planned to estimate the probabilities of transmission by the various routes. For example, double layers of wire screening between pens, with sufficient distance between the layers to prohibit direct

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74 contact, could be used to assess the probability of aerosol transmission. Pen designs in which water containers are shared, but direct contact and aerial transmission are precluded could generate data addressing the role of transmission via water. Tortoises' strength, determination, and digging abilities must all be considered when designing such experiments. Seropositive, clinically healthy, culture and PCR negative individuals may be suitable for relocation, restocking, and repatriation programs, but clinically ill animals should not be used in such efforts. However, the proportion of tortoises that appear to have recovered from disease and eliminated the mycoplasma, while actually harboring the organism, is unknown. As evidenced in the preceding chapter, some of those tortoises may have latent infections that will recrudesce under stressful conditions.

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CHAPTER 6 VERTICAL TRANSMISSION OF MYCOPLASMA AGASSIZII Introduction Many mycoplasmas can be transferred vertically, that is, from mother to offspring in utero or in ovo (see Chapter 1). Before appropriate decisions can be made concerning the fate of desert or gopher tortoises infected with M. agassizii, particularly those from threatened or endangered populations, questions regarding the vertical transmissibihty of M. agassizii need to be resolved. If M. agassizii is not transmissible via eggs, the options for preserving genetic material are increased. I designed experiments to test the hypotheses that M. agassizii is transferred in ovo to gopher tortoise hatchlings and that levels of specific antibodies against M. agassizii in egg yolk and hatchling plasma are associated with those found in maternal plasma. Methods Adult tortoises were housed as detailed in Chapter 2, and selected as specified in Chapter 5. In order to obtain clutches of eggs, pairs were established in August, 1994, as detailed in Chapter 5. The sampling schedule was as detailed in Chapter 5. 75

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76 Egg Collection and Incubation Egg development was monitored by radiography (6 mas, 62-76 kv) beginning in April of 1995 and 1996. When eggshells were judged to be calcified adequately, oviposition was induced by intravenous injection of arginine vasotocin (Sigma) at approximately 10-12 picograms/kg. Tortoises were monitored until clutch deposition was complete or for a minimum of 4 hr. Eggs were collected as they were laid and placed in a sanitized container (washed with 1:10 bleach solution and air-dried upside down) partially filled with sterilized vermiculite moistened with an equal weight of sterilized water. Female identification number and letter indicating order of deposition were written on each egg with a graphite pencil swabbed with 70% ethanol. After approximately half the clutch was laid, cloacal swabs for mycoplasma culture were taken, streaked onto SP4 agar, placed in SP4 broth, and incubated as described previously. In 1995, eggs were incubated at 29°C until hatching. In 1996, eggs were incubated at 27, 29 or 3 1°C. Approximately 1 wk before hatching each egg was cleansed of adhering venniculite with clean gauze and placed into a sanitized plastic container. After hatching, resorption of yolk, and closing of the umbilicus, hatchlings were placed in containers with clean sand. Hatchlings were maintained separately from adults, at ambient temperature and light cycles, and fed natural foods supplemented with commercially available vegetables until release into outdoor pens, where they were fed natural foods. Culture and PCR Procedures Eggs were taken at various times during incubation for mycoplasma culture and antibody detection. At pipping, chorioallantoic-amniotic fluid (CAF) was collected and

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77 frozen for later analyses. For culture, 100 ul samples of yolk and albumin, and small pieces of membrane were added to 900 ul of SP4 broth. After 48 hr incubation at 30°C, 500 ul of culture were removed for PCR analysis. All CAF samples obtained at pipping were treated in the same manner. In 1996, one or two pooled samples consisting of 100 ul of CAF from each of 3 6 eggs, mixed well, were made for each clutch and processed as described. ELISA Procedures Based on preliminary experiments using several methods of antibody extraction, the supernatant resulting from mixing 1 ml of yolk with 1 ml PBS-AZ provided the most efficient fraction for detection of antibodies (G. S. McLaughlin and L M. Schumacher, unpub. data). Blood samples for ELISA were collected by cardiocentesis from most hatchlings at approximately 2-4 wk of age. In 1995, the goal was to obtain a blood sample from each hatchling. In 1996, sampling effort was reduced to a maximum of five samples per clutch due to the low coefficients of variation for the 1995 samples. In 1995, hatchling ELISA samples were run as previously described except that three dilutions from the range 1 : 1 to 1 : 8 were run in duplicate, depending on the sample volume obtained. Sample dilutions were based on previous experiments with plasma from desert tortoise hatchlings (I. M. Schumacher, unpub. data). All hatchling or egg samples were run on the same plate as 1994 and 1995 samples from the corresponding female and the presumed sire, if possible. If samples were split between two plates, plates were run on the same day with the same reagents and positive and negative controls. In 1996, samples were run in triplicate at a dilution of 1:2, as that was the dilution used for analysis

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78 of the 1995 samples. Additionally, that modification reduced the necessary blood volume from approximately 400 |^1 to 150 ul, reducing stress to the hatchlings. Samples collected in 1995 and 1996 from the corresponding female were run on the same plate as samples from their eggs and/or hatchlings. Results Clutch Sizes. Fertility and Hatching Rates Twenty-six clutches, 13 each year, were collected. Arginine vasotocin was used to induce oviposition of 20 clutches and the rermining six clutches (five in 1995 and one in 1996) were laid in the pens. In 1995, 13 tortoises developed 115 eggs (mean = 8.8), and 103 eggs were recovered (Table 6-1). Five females laid entire clutches in the pens, four of which were removed within 1 wk of laying. One female laid nine eggs in an inaccessible area of her pen. Two females that laid incomplete clutches on induction laid the remaining eggs (n = 3) in their pens. Twentyfive eggs were removed for culture, PCR, and ELISA samples. Of those, five were fertile and removed while viable, four had died, and five were infertile. I could not determine fertility on the remaining 1 1. Overall clutch sizes (Table 6-1) were larger in 1996 (mean = 11.1) than in 1995 (mean = 8.8, 2-tail /-test = 2.78, 17 df P = 0.013), as were those of seronegative females (1 1.7 vs 9.8, 2-tail Mest = 2.45, 9 dfi P = 0.037). In 1996, clutch sizes of seronegative females (mean = 11.7) were larger than those of seropositive females (mean = 10.3, 2-tail Mest = 2.30, 1 1 df P = 0.042). Thirteen females developed 144 eggs, 130 of which were recovered. One female laid 1 1 eggs that were incompletely calcified and broke during the

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79 laying process. Five eggs were laid in the pen and six while in transport; a usable sample for culture and PCR was obtained from only one egg. Three females retained one egg each, the fates of which were unknown. One fertile egg was donated for the purpose of establishing embryonic tortoise cell lines, and 19 eggs were sampled. Of those, 13 eggs were infertile, four had died during incubation, and I could not determine fertility of two. In 1995, 78 eggs pipped and 77 hatchlings were produced in the laboratory (Table 6-1), and nine hatchlings were recovered from the 12 eggs laid in the pens (1/1, 1/2, 7/9). The tortoise that pipped but did not hatch had an abnormal yolk sac; it was either malformed or had ruptured, and yolk was found throughout the coelom The hatchling was in respiratory distress (cyanotic and mouth breathing), and was euthanatised. Another fetus appeared to be full term, but had spina bifida in the cervical region, and only one kidney. In 1996, 1 1 1 eggs produced 1 13 viable hatchlings (Table 6-1). One egg produced one healthy hatchling and one that was anencephalic. Two other eggs, from a different clutch, each produced healthy twins. There were no differences in fertility or hatching rates between infected and uninfected females for either year individually or both years combined (x 2 tests, 1 df all P > 0.75). Fertility and hatching rates did not differ significantly between years (% 2 tests, 1 df, all P > 0.20). Culture and PCR Results No cultures or PCR assays of cloacal samples, egg yolk, albumin, or membranes were positive for mycoplasma, and no hatchlings isolated from adults developed clinical signs (see Chapter 5 for evidence of horizontal transmission from adults to hatchlings).

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Table 6-1. Clutch data from gopher tortoise upper respiratory tract disease vertical ransmission study, 1995 and 1996. Serostatus 1995 1996 negative positive total negative positive total No. clutches 8 5 13 7 6 13 No. eggs 78 37 115 82 62 144 Mean a 9.8 7.4 a 8.8 a,b 11.7 b 10.3 a 11.1 No. recovered 75 28 103 69 61 130 No. sampled 16 9 25 11 8 19 Fertility rates 94% 97% 95% 88% 92% 90% No. hatchlings 61 26 87 52 61 113 Hatching rates 90% 93% 91% 86% 88% 87% CAP samples 59 19 78 51 51 102 Blood samples 51 25 76 29 28 57 Clutch sizes differed significantly between 1995 and 1996 for seronegative tortoises (2-tail Mest = 2.45, P = 0.037), and for overall clutch size (2-tail Mest = 2.78, P = 0.013). b Clutch sizes differed significantly between seronegative and seropositive tortoises for 1996 (2-tail Mest = 2.30, P = 0.042). ELISA Results For ELISAs in 1995, 76 hatchlings and 34 yolks were sampled for a total of 96% of all available eggs and hatchlings sampled. Total sampling effort per clutch ranged from 75-100%, with 50-100% of hatchlings sampled. In 1996, 57 hatchlings and 19 yolks were sampled, for a total sampling effort of 58%, with sampling effort per clutch of 42-100%. ELISA values in egg yolks and hatchlings were correlated with maternal ELISA values in Autumn of the prior calendar year (r = 0.68, R 2 = 0.461, P < 0.0002, n = 25; Figure 6-1), indicating that specific antibodies were transferred via the egg yolks to the hatchlings.

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82 Discussion Unlike various mycoplasmal infections of rodents and poultry (Simecka et al. 1992) and wild ducks (Goldberg et al. 1995), there was no evidence to support a hypothesis of vertical transmission of M. agassizii. Therefore, it should be possible to collect eggs from infected female tortoises, incubate the eggs, and release the hatchlings with no risk of spreading the disease. Vitellogenesis begins in August or September and continues to December (Taylor 1982), providing an extensive period for deposition of antibodies. In spite of that, the antibody level in egg yolks and hatchling plasma was only 10 -20% of that in the maternal plasma. Thus, there is no evidence that female gopher tortoises are sequestering antibodies in the egg components. This is in contrast to birds, particularly chickens, which deposit large amounts of antibodies in their eggs in a very short time period. Antibody levels in chicken eggs can be several orders of magnitude greater than that in the maternal plasma. Hatchling antibody levels decline slowly over the first year of life, as the maternal antibodies are broken down (I. M. Schumacher, unpub. data). This process is much slower in tortoises than in mammals and birds, where such passively acquired antibodies decline within weeks or a few months after birth or hatching. It is not known if antibodies against M. agassizii affect juvenile responses to infection with the organism Although adult tortoises that have developed an antibody response against M. agassizii respond very quickly and adversely to subsequent exposure (Chapter 4), that response may be mediated more by cellular immune components (e.g., macrophages and heterophils) than by

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83 humoral. Hatchling and juvenile tortoises with maternal antibodies may be protected from some effects of infection if the antibodies are neutralizing in nature, and if they can be mobilized appropriately against the mycoplasma. Research needs to be conducted to address these areas.

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CHAPTER 7 ENVIRONMENTAL TRANSMISSION OF MYCOPLASMA AGASSIZII Introduction Some mycoplasmas can remain viable for at least several days in the environment (Chandiramani et al. 1966), and for many years under refrigeration or freezing (Yoder and Hofstad 1964). One method of controlling mycoplasma infections in domestic stock is to depopulate a farm or facility, spray buildings and equipment with disinfectants, wait an appropriate amount of time, and then reintroduce uninfected stock (Anonymous 1989). In order to determine appropriate time frames for restocking tortoises, knowing the length of time Mycoplasma agassizii remains viable in the environment is critical. Unfortunately, direct viability testing is extremely difficult and impractical, if not impossible. Many soil bacteria and fungi in the tortoise burrow and surrounding environment grow very rapidly in culture and quickly overtake any mycoplasma colonies that might be present. However, the risk of environmental transmission of M. agassizii is an important parameter in the decision making process. In order to address this question, I designed an experiment to test the hypothesis that environmental transmission of M. agassizii would occur in tortoises introduced to pens previously occupied by infected, clinically ill tortoises. 84

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85 Methods As a result of previous and concurrent experiments, 13 seropositive tortoises infected with M. agassizii and 15 seronegative tortoises with no clinical signs and negative culture and PCR results were available. The 13 positive tortoises were captured and reinoculated with approximately 10 8 colony forming units (CFU) ofM agassizii to ensure active clinical disease and shedding of bacteria at the time scheduled for introducing healthy tortoises to the pens (see Chapter 4). Six weeks following inoculation, the 13 tortoises were captured, as were the 1 5 healthy tortoises. Nasal flush samples were collected from each infected tortoise to analyze by PCR Immediately after capture and removal of five infected tortoises from their pens, a healthy tortoise was put into the pen. Transfers were carried out within 15 min. Transfers were also carried out at 3 (n = 4), 7 (n = 3), and 10 (n = 3) days following removal of the infected tortoises. Because only 13 ill tortoise pens were available, one of the day 7 and one of the day 10 transfers consisted of pairs of tortoises. Eight weeks following transfer, the 15 tortoises were captured, assessed clinically, photographed, and blood and nasal flush samples were collected. Samples were handled as previously described. Results All infected tortoises were clinically ill at the time of capture, although PCR results for all animals were negative. Transferred tortoises explored their new surroundings and usually entered the burrows within 1 hr, and some entered within 15 min. At 8 wk posttransfer, no transfer tortoise was clinically ill, or had positive culture or PCR results. No tortoise showed an increase in ELISA values indicative of possible infection (Table 7. 1).

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86 Table 7.1. ELISA results from initially clinically healthy, culture and PCR negative tortoises transferred into pens previously occupied by clinically ill, culture or PCR positive tortoises. Ratios for equal ELISA values may differ due to plate--to~plate variation in the Tortoise Pre-transfer ELISA Post-transfer ELISA Transfer ID Value Ratio Value Ratio Day 1 AO 0.275 1.89 0.242 1.66 n \j 129 0.164 1.17 0.137 0.98 0 160 0.185 1.75 0.236 1.80 0 201 0.217 1.70 0.192 1.51 0 205 0.471 3.37 0.409 2.93 0 110 0.119 0.94 0.137 1.08 3 211 0.222 2.00 0.234 2.11 3 219 0.352 2.72 0.184 1.43 3 311 0.134 1.21 0.144 1.29 3 159 0.278 2.12 0.160 1.23 7 226 0.279 2.63 0.212 2.00 7 261 0.378 2.92 0.287 2.21 7 119 0.113 1.02 0.120 1.08 10 125 0.063 0.60 0.054 0.52 10 140 0.134 1.23 0.152 1.40 10 Discussion Although the PCR results for the re-infected tortoises were negative at the time of capture for transfer, the PCR is limited in its sensitivity. The lower limit of detection is approximately 1000 CFU (D. R Brown, unpub. data), whereas the infectious dose of M agassizii may be 10 or fewer CFU (Brown et al. 1996b). Therefore, tortoises could have been shedding infective doses of bacteria and still have negative PCR results. In other studies, 100% of 28 naive tortoises seroconverted by 8 wk PI, with all tortoises tested at 6 wk PI having significantly increased antibody levels (Brown, D. R, 1996b; Schumacher, unpub. data; Chapter 4). No tortoises in this study had increased

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antibody levels, and it is unlikely that insufficient time had passed for the tortoises to develop an immune response. Environmental transmission of M. gallisepticum is of considerable concern to the poultry industry, necessitating disinfection of premises and equipment, and a 2-6 wk fallow period before introducing new stock (Anonymous 1989, M. B. Brown pers. comm.). The M gallisepticum strain causing conjunctivitis in house finches also can survive in the environment and cause infections in individuals later housed in the same facility (Ley et al. 1996, Luttrell et al. 1996). Environmental transmission of M. agassizii in the wild may not be of great concern. However, equipment used for capturing, handling, holding, and transporting tortoises should be cleaned after each use by spraying or wiping with bleach, ethanol or other disinfectant solution. Care should be taken to dispose properly of all gloves or drapes that may have become contaminated. Clinically ill tortoises should not be housed in direct contact with other animals, nor should indirect contact be allowed. Infected tortoises should not be able to sneeze on other tortoises, nor should water or food dishes be shared between pens without disinfection. Because the sample sizes at each time point were very small, further research, with more rigorous attempts to isolate mycoplasmas and quantify shedding, needs to be conducted. For the purposes of relocation, restocking, or repatriation efforts, a short fallow period, perhaps 2 wk, is probably sufficient to prevent environmental transmission of M. agassizii.

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CHAPTER 8 CONSERVATION AND MANAGEMENT IMPLICATIONS OF UPPER RESPIRATORY TRACT DISEASE The research reported in this dissertation has built on and extended the findings of research on upper respiratory tract disease in desert and gopher tortoises. Based on the knowledge that Mycoplasma agassizii causes clinical signs and lesions of URTD and elicits an antibody response detectable by an ELISA, experiments were designed to compare the pathological effects of the disease in gopher tortoises to those in desert tortoises, investigate secondary immune responses, and determine routes of transmission. The knowledge gained can be applied to conservation and management decisions, and new areas of research have been suggested. Implications for Conservation and Management When tortoises are impacted by development, mining, agriculture, or forestry practices, decisions must be made regarding their disposition. Tortoises can be ignored, temporarily removed and confined to pens for later return to the site, permanently moved to a currently inhabited site, moved to a formerly occupied site, donated to research or educational facilities, or euthanatised. The choices open for a particular population of tortoises depend on the location, historical, current, and future site use, surrounding land 88

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89 use patterns, importance of the population in maintaining genetic variability, and political and social factors. No one prescription will be ideal for all situations, and it is difficult, if not impossible, to develop a set of prescriptions that will cover all permutations of the above factors. However, guidelines for making decisions can be developed, and some are presented in this chapter. Establishing Goals The goals of the management action(s) must be established before decisions can be made regarding what data to collect, or how to design survey or monitoring programs. Management tactics designed to establish, create, or maintain a URTD-free population will differ from those intended to maintain the status quo relative to disease agents. If the genetic material represented by tortoises in defined or isolated populations (e.g., those in South Carolina, Mississippi, or Louisiana, and some in Alabama) is important for conservation purposes, aggressive efforts to protect and maintain the gene pool may be of primary concern. Understanding URTD and Test Results An understanding of test results is necessary for proper interpretation and application. For field personnel, recognition of clinical signs of URTD is an important skill to gain. Differentiation of signs of URTD from other conditions is difficult even for experts, as the signs are nonspecific, but some conditions can be recognized fairly easily (e.g., foreign body in or trauma to the eye causing discharge, or nasal "discharge" due to

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90 drinking or eating). Tortoise behavior interacts with clinical disease to affect transmission probabilities, so understanding behavior also is important in the decision making process. A clinically healthy tortoise, with negative ELISA, culture, and PCR results is probably free from URTD. A positive ELISA, in the absence of clinical illness and positive culture or PCR results, indicates only that the tortoise has been exposed to M. agassizi. Because clinical signs and culture and PCR results can vary over time, we cannot predict if or when a seropositive tortoise will break with clinical disease and begin shedding bacteria. The more stress to which an animal is subject, such as human intrusion, handling or transport, drought or other extreme weather conditions, the more likely it is to have a disease recurrence. Repeated exposures to M. agassizii elicit more intense immunological responses by the tortoises, potentially leading to autoimmune responses that may contribute to the more severe lesions seen in longer-term infections, as well as possible liver pathology. Tortoises with rapid responses to the mycoplasma, with copious mucus production, may be more likely to transmit the agent to conspecifics, as their energy reserves have not been depleted by the disease process. A clinically ill tortoise with positive culture or PCR results, regardless of ELISA results, probably is capable of transmitting M agassizii. The more active a tortoise is, and the greater its daily movements, the more likely it is to spread the bacteria through a colony and foment an outbreak of URTD. Behaviorally, male tortoises have larger home ranges and more intraspecific contacts, so are probably at greater risk of coming in contact with and spreading the pathogen.

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91 A clinically healthy tortoise (i.e., one with no nasal or ocular discharge or other signs) may have positive culture and PCR results, with either positive or negative ELISA results. It may be in the early stages of infection or recrudescence. In the former case, the ELISA value may be negative, but should rise within 6 wk. In the latter case, the ELISA value may be quite high. Such tortoises may be capable of transmitting the mycoplasma under the appropriate conditions. Although direct contact (nose-to-nose) seems to be the most important route, transmission through water or on food cannot be ruled out. When tortoises drink, they often expel water through their noses for short distances, up to 50 cm, or sneeze forcefully after drinking, aerosolizing the contents of the nasal passages. If tortoises are in close contact with one another, spatially or temporally, such occurrences may allow transmission of M. agassizii. Tortoises with slight nasal discharge, virtually undetectable, can also aerosolize bacteria by sneezing. Although long-term studies on the effect of URTD on survival of individuals and populations have not been conducted, the evidence from surveys of desert tortoise populations and from Sanibel Island indicate that the disease can have severe negative impacts on population viability. Declines of 25-50% over 1-3 yr, and of 30-90% over 10 yr, are catastrophic for species that take 10-20 yr to reach maturity and have recruitment rates of 1-2%. Without marked improvement in recruitment rates, affected populations are unlikely to recover within a reasonable time frame. Developing Questions and Conducting Surveys or Monitoring Programs Third, questions related to management goals, and taking into account test interpretation and tortoise behavior, must be developed. Some of the most common

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92 questions are: 1) What is the prevalence of antibodies against M. agassizii in the population of interest? 2) What is the distribution of seropositive tortoises relative to gender, age, habitat type, geography, or land use patterns? 3) Has the seroprevalence changed over time? 4) Is clinical disease present on the sites? 5) IsM agassizii detectable from any tortoises? 6) Is there evidence, either from demographic profiles or tortoise remains, of large die-offs, or gaps in recruitment? Once the goals have been established and questions developed, surveys or monitoring programs can be designed to collect the necessary samples and data. After field work has been conducted, samples collected and analysed, and test results analysed, the information can be used to develop management programs or research questions. When trapping tortoises for studies or relocation, it is important to note any clinical signs that are present. Simply being aware of the possibility that URTD exists on a site may make field personnel more likely to detect clinical cases. While this does not guarantee that every case of URTD will be caught, it could give an early warning of potential problems, prevent sick tortoises from being put in holding pens with healthy ones, or transmitting the agent on equipment. It is easy to spray equipment with a mild bleach solution, or wipe down calipers and other equipment with alcohol soaked gauze. It is more difficult, and expensive, to construct separate holding facilities for healthy and sick tortoises, but alternatives are available. Ill tortoises can be kept in containers for a few days, long enough for monitoring and until test results come back.

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93 Weighing Management Options and Formulating Management Plans Knowing that URTD exists or has existed on a site, and the distribution of seropositive tortoises, may suggest management strategies to cope with potential problems. If tortoises testing positive are concentrated in one area, there may be opportunities to improve habitat so tortoises are not as stressed in times of food shortage, drought, or other adverse conditions. More monitoring time could be allocated to such areas in order to get an early warning in the case of a disease outbreak. If the site is close to or at carrying capacity, the population could be reduced so the remaining tortoises are under less intraspecific stress. Eggs could be collected from that site to use in headstart programs to ensure the genes are not lost in case of an outbreak. If there is a population of tortoises into which relocated ones will be released, and the goal is to maintain the status quo relative to disease agents, then both populations need to be tested. If the recipient population and the donor population have approximately the same level of seropositive tortoises, and the site is well below carrying capacity, then it is probably acceptable to release seropositive tortoises, as long as the overall rate is not increased, and no tortoises actively shedding bacteria are released. Releasing clinically ill tortoises, regardless of ELISA results, is not recommended. If a tortoise has a nasal discharge, palpebral edema, and conjunctivitis that persist for more than 24 hr, there is a substantial risk that the tortoise is shedding mycoplasma. Tortoises may shed more bacteria early in the infection, before they have developed a strong antibody response. Due to the secondary immune responses (Chapter 4), seropositive

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94 tortoises that may have cleared the bacteria would be at risk of developing severe disease when exposed to a clinically ill tortoise, decreasing their survival probability. If the goal is to establish, create, or maintain a URTD-free population, the question remains of what to do with seropositive, clinically healthy, culture and PCR negative individuals. If there is no secure place to hold the animals or a site where they can be released, then the most acceptable choice may be euthanasia. Euthanasia may save the animals from a long slow death, and prevents the chance of exposing negative animals. Alternatively, intensive monitoring of the population, with special attention paid to seropositive tortoises and samples collected several times per year, may allow rapid detection and removal of ill animals. However, such careful monitoring is beyond the capacity of most management agencies, and annual or biennial sampling may be all that can be accomplished. Researchers sometimes need seropositive animals for projects. If one can be found, it is possible that animals could be donated. However, there are many issues to consider with that approach, mostly for the researcher. Researchers usually cannot accept animals that are made available suddenly, particularly for studies that require animals to be kept in laboratory settings. Costs, objectives, confounding factors of length of disease, genetics and site of origin, are all issues that must be addressed. It is expensive to keep tortoises in captive research settings, and safeguards must be in place to prevent their escape or theft. Research can be a politically and socially sensitive subject due to potentials for pain, illness, and death of animals, and nearly all animals are euthanatised at the end of the studies.

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95 If the facilities and personnel are available, and the genetic material represented by the sick or seropositive tortoises is important, then a captive breeding program may provide the best solution. Clinically ill animals or those that test positive by culture or PCR should be maintained separately from clinically healthy, culture and PCR negative animals. Eggs can be collected from the females by induction of oviposition or at natural laying, or nests can be dug up. Eggs should not be allowed to hatch and hatchlings emerge from nests in pens inhabited by ill adults. Hatchlings can contract URTD from adults, and they generally become very ill very quickly. Eggs can be artificially or naturally incubated, and hatchlings released immediately or headstarted. Summary of Conservation and Management Implications In summary, the following points must be considered when making conservation and management decisions relative to URTD in gopher tortoises: 1) Goals must be clearly established. 2) Personnel must have appropriate training to recognise URTD, collect necessary samples, and interpret results. 3) Clear questions must be formulated. 4) Survey and monitoring programs must be developed and implemented, and precautions taken to ensure detection and prevent spread of URTD. 5) Management options must be weighed, and plans formulated and implemented that are consistent with established goals. a) Habitat manipulations. b) Relocate tortoises and maintain status quo relative to URTD.

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96 c) Establish, create, or maintain URTD-free populations by careful monitoring, testing, and removal of infected tortoises. d) Donate animals to research projects. e) Establish captive breeding programs. f) Euthanasia. Further Research It is not known how long Mycoplasma agassizii has been in gopher and desert tortoises, or if it occurs naturally in tortoises in other parts of the world. At least one other mycoplasma that causes URTD has been found in some populations of desert tortoises, and there may be other etiologic agents. An iridovirus was found in an ill gopher tortoise from Sanibel Island, and may have been the cause of the disease in that animal (Westhouse et al. 1996). Further research needs to be conducted on that agent. The long-term effects of M. agassizii infection need to be studied. There are indications that liver function might be affected (Chapter 3), and that reproductive physiology is altered (Rostal et al. 1996). The impacts of altered basking and foraging behavior need to be quantified, as do the rates of transmission within natural populations. The role of maternal antibodies on the course of disease in hatchlings and juveniles is not understood. In order to fully understand the impacts of URTD on tortoise populations, many ares of research need to addressed, surveys undertaken, and long-term monitoring programs established.

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103 Kerr, K. M., and N. O. Olson. 1967. Pathology in chickens experimentally inoculated or contact-infected with Mycoplasma gallisepticum. Avian Diseases 1 1 : 559-578. Kirchhoff, H., R. Schmidt, H. Lehmann, H. W. Clark, and A. C. Hill. 1996. Mycoplasma elephantis sp. nov., a new species from elephants. International Journal of Systematic Bacteriology 46: 437-441. Kleven, S. H., H. H. Fan, and K. Turner. 1996. Displacement of virulent Mycoplasma gallisepticum in chickens with live vaccines. IOM Letters 4: 284-285. Knowles, C. 1989. A survey for diseased desert tortoises in and near the Desert Tortoise Natural Area, Spring 1989. Report prepared for the Bureau of Land Management, Riverside, California, Contract No. CA 950-(T9-23), 26 pp. Krause, D. C, and D. Taylor-Robinson. 1992. Mycoplasmas which infect humans. Pp. 417-444 in: Maniloff, J., R. N. McElhaney, L. R. Finch, J. B. Baseman (eds.). Mycoplasmas: molecular biology and pathogenesis. American Society for Microbiology, Washington, D. C. Kume, K\, Y. Kawakubo, C. Morita, E. Hayatsu, and M. Yoshioka. 1977. Experimentally induced synovitis of chickens with Mycoplasma synoviae: effects of bursectomy and thymectomy on course of infection for the first four weeks. American Journal of Veterinary Research 38: 1595-1600. Lai, W. C, M. Bennett, and S. P. Pakes. 1996. Successful immunization of mice after infection with virulent Mycoplasma pulmonis. IOM Letters 4: 52. Landers, J. L., and D. W. Speake. 1980. Management needs of sandhill reptiles in southern Georgia. Proceedings of the Annual Conference Southeastern Association of Fish and Wildlife Agencies 34: 515-529. Landers, J. L., J. A. Garner, and W. A. McRae. 1980. Reproduction of the gopher tortoise (Gopherus polyphemus) in southwestern Georgia. Herpetologica 36:353-361. Landers, J. L., W. A. McRae, and J. A. Garner. 1982. Growth and maturity of the gopher tortoise in southwestern Georgia. Bulletin of the Florida State Museum, Biological Sciences 27:81-1 10. Lawrence, K., and J. R Needham. 1985. Rhinitis in long-term captive Mediterranean tortoises (Testudo graeca and T. hermanni). Veterinary Record 1 17: 662-664. LevelL J. P. 1995. A field guide to reptiles and the law. Serpent's Tale, Excelsior, MN. 240 pp.

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Ley, D. H., J. E. BerkhofF, K. Joyner, L. Powers, and S. Levisohn. 1996. Molecular epidemiology investigations of Mycoplasma gallisepticum conjunctivitis in songbirds by random amplified polymorphic DNA (RAPD) analysis. IOM Letters 4: 53-54. Ley, D. H., J. E. Berkhoff, and J. M. McLaren. 1996. Mycoplasma gallisepticum isolated from house finches {Carpodacus mexicanus) with conjunctivitis. Avian Diseases 40: 480-483. Lindsey, J. R, H. J. Baker, R G. Overreach, G. H. CasselL and C. E. Hunt. 1971. Murine chronic respiratory tract disease: Significance as a research complication and experimental production with Mycoplasma pulmonis. American Journal of Pathology 64: 675-708. Linley, T. A., and H. R. Mushinsky. 1994. Organic composition and energy content of eggs and hatchlings of the gopher tortoise. Pp. 1 13128 in R. B. Bury and D. J. Germano (eds.). Biology of North American tortoises. National Biological Survey, Fish and Wildlife Research 13. Lips, K. R 1991. Vertebrates associated with tortoise {Gopherus polyphemus) burrows in four habitats in south-central Florida. Journal of Herpetology 25: 477-481. Lohoefener, R, and L. Lohmeier. 1981. Comparison of gopher tortoise {Gopherus polyphemus) habitats in young slash pine and old longleaf pine areas of southern Mississippi. Journal of Herpetology 15:239-242. LuttrelL M. P., S. H. Kleven, and W. R Davidson. 1991. An investigation of the persistence of Mycoplasma gallisepticum in an eastern population of wild turkeys. Journal of Wildlife Diseases 27: 74-80. LuttrelL M. P., J. R Fischer, D. E. Stallknecht, and S. H. Kleven. 1996. Field investigation of Mycoplasma gallisepticum infections in house finches {Carpodacus mexicanus) from Maryland and Georgia. Avian Diseases 40: 335-341. LyelL A., A. M. Gordon, H. M. Dick, and R. G. Sommerville. 1967. Mycoplasmas and erythema multiforme. Lancet 2: 1116-1118. Macdonald, L. A., and H. R. Mushinsky. 1988. Foraging ecology of the gopher tortoise, Gopherus polyphemus, in a sandhill habitat. Herpetologica 44:345-353. Mann, T. M. 1990. The status of Gopherus polyphemus in South Carolina. Pp. 88-130 in C. K Dodd, Jr., R E. Ashton, Jr., R Franz, and E. Wester (eds.). Proceedings of the 8th Annual Meeting Gopher Tortoise Council, Florida Museum of Natural History, Gainesville, FL. ManseL J. K., E. C. Rosenow HI, T. F. Smith, and J. W. Martin, Jr. 1989. Mycoplasma pneumoniae pneumonia. Chest 95: 639-646.

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105 Markham, J. F., P. C. Scott, and K. G. Whithear. 1996. Field and laboratory studies on a live attenuated Mycoplasma synoviae vaccine. IOM Letters 4: 286. McLaughlin, G. S. 1990. Ecology of gopher tortoises (Gopherus polyphemus) on Sanibel Island, Florida. M. S. Thesis. Iowa State Univ., Ames. 124 pp. McRae, W. A., J. L. Landers, and J. A. Garner. 1981. Movement patterns and home range of the gopher tortoise. American Midland Naturalist 106: 165-179. Mohan, K., C. M. Foggin, P. Muvavarirwa, J. Honywill, and A. Pawandiwa. 1995. Mycoplasmaassociated polyarthritis in farmed crocodiles (Crocodylus niloticus) in Zimbabwe. Onderstepoort Journal of Veterinary Research 62: 45-49. Mohan, K., C. M. Foggin, P. Muvavarirwa, and J. Honywill. 1996. Experimental trial with an alum-precipitated vaccine against Mycoplasma-nssocistQd polyarthritis in farmed crocodiles (Crocodylus niloticus). IOM Letters 4: 287. Murray, H. W., H. Masur, L. B. Senterfit, and R. B. Roberts. 1975. The protean manifestations of Mycoplasma pneumoniae infections in adults. American Journal of Medicine 58: 229-242. Mushinsky, H. R 1985. Fire and the Florida sandhill herpetofaunal community: with special attention to responses of Cnemidophorous sexlineatus. Herpetologica 4 1 : 333-342. Mushinsky, H. R. 1986. Fire, vegetation structure, and herpetofaunal communities. Pp. 383-388 in Z. Rocek (ed.). Studies in herpetology. Charles University Press, Prague. Mushinsky, H. R, and D. J. Gibson. 1991. The influence of fire on habitat structure. Pp. 237-259 in S. S. Bell, E. D. McCoy, and H. R. Mushinsky (eds.). The physical arrangement of objects in space. Chapman and Hall, London. Mushinsky, H. R, and E. D. McCoy. 1994. Comparison of gopher tortoise populations on islands and on the mainland in Florida. Pp. 1 13-128 in R B. Bury and D. J. Germano (eds.). Biology of North American tortoises. National Biological Survey, Fish and Wildlife Research 13. Mushinsky, H. R, D. S. Wilson, and E. D. McCoy. 1994. Growth and sexual dimorphism of Gopherus polyphemus in central Florida. Herpetologica 50: 1 19-128. Naftalin, J. M., G. Wellish, Z. Kanana, and D. Diengott. 1974. Mycoplasma pneumoniae septicemia. Journal of the American Medical Association 228: 565. Nettles, V. 1996. Reemerging and emerging infectious diseases: Economic and other impacts on wildlife. ASM News 62: 589-591.

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106 Nunoya, T., M. Tajima, T. Yagihashi, and S. Sannai. 1987. Evaluation of respiratory lesions in chickens induced by Mycoplasma gallisepticum. Nippon Juigaku Zasshi 49: 621-629. Oboegbulem, S. I. 1981. Enzootic pneumonia of pigs: a review. Bulletin of Animal Health and Production in Africa 29: 269-274. OftedaL O. T., T. E. Christopher, and M. E. Allen. Upper respiratory tract disease in hatchling tortoises. Proceedings of a Conference on Health Profiles, Reference Intervals, and Diseases of Desert Tortoises, October 31 November 3, 1996, Soda Springs, CA. Pp. A-44 A-47. Paling, R. W., S. Waghela, K J. Macowan, B. R Heath. 1988. The occurrence of infectious diseases in mixed farming of domesticated wild herbivores and livestock in Kenya. EL Bacterial diseases. Journal of Wildlife Diseases 24: 308-316. Poveda, J. B., J. GiebeL H. Kirchhoff, A. Fernandez. 1990. Isolation of mycoplasmas from a buzzard, falcons and vultures. Avian Pathology 19: 779-783. Poveda J. B., J. Giebel, J. Flossdorf, J. Meier, H. Kirchhoff. 1994. Mycoplasma buteonis sp. nov., Mycoplasma falconis sp. nov., and Mycoplasma gypis sp. nov., three species from birds of prey. International Journal of Systematic Bacteriology 44: 94-98. Rocke, T. E., and T. M. Yuill. 1987. Microbial infections in a declining wild turkey population in Texas. Journal of Wildlife Management. 51: 778-782. Rocke, T. E., T. M. Yuill, and T. E. Amundsen. 1988. Experimental Mycoplasma gallisepticum infection in captive reared wild turkeys. Journal of Wildlife Diseases 24: 528-532. Rosenbusch, R. F. 1987. Immune responses to Mycoplasma bovoculi conjunctivitis. Israeli Journal of Medical Science 23: 628-631. RostaL D. C, V. A. Lance, J. S. Grumbles, and I. M. Schumacher. 1996. Effects of acute respiratory tract disease on reproduction in the desert tortoise, Gopherus agassizii: hormones, egg production and hatchling success. Proceedings of a Conference on Health Profiles, Reference Intervals, and Diseases of Desert Tortoises, October 31 November 3, 1996, Soda Springs, CA. Pp. A-48 A-52. Samuel, M. D, D. R Goldberg, C. B. Thomas, and P. Sharp. 1995. Effects of Mycoplasma anatis and cold stress on hatching success and growth of mallard ducklings. Journal of Wildlife Diseases 31: 1 721 78. SAS Institute Inc. 1988. SAS/STAT™ users guide, release 6.03 edition. Cary, NC: SAS Institute Inc. 1028 pp.

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Schumacher, I. M, M. B. Brown, E. R. Jacobson, B. R. Collins, and P. A. Klein. 1993. Detection of antibodies to a pathogenic mycoplasma in desert tortoises (Gopherus agassizii) with upper respiratory tract disease. Journal of Clinical Microbiology, 3 1 : 1454-1460. Simecka, J. W., J. K. Davis, M. K. Davidson, S. E. Ross, C. T. K. H. Stadtlander, and G. H. Cassell. 1992. Mycoplasmas which infect animals. Pp. 391-415 in: Maniloff, J., R N. McElhaney, L. R Finch, J. B. Baseman, eds. Mycoplasmas: molecular biology and pathogenesis. American Society for Microbiology, Washington, D. C. Snipes, K. P., and E. L. Biberstein. 1982. Pasteurella testudinis sp. nov.: a parasite of desert tortoises. International Journal of Systematic Bacteriology 32: 201-210. Springer, W. T., C. Luskus, and S. S. Pourciau. 1974. Infectious bronchitis and mixed infections of Mycoplasma synoviae and Escherichia coli in gnotobiotic chickens. I. Synergistic role in the airsacculitis syndrome. Infection and Immunity 10: 578-589. Stalheim, O. H. 1983. Mycoplasmal respiratory diseases of raminants: a review and update. Journal of the American Veterinary Medical Association 182: 403-406. Stipkovits, L. 1979. The pathogenicity of avian mycoplasmas. Zentralblatt fur Bakteriologie, Parasitenkunde Infektionskrankheiten und Hyggiene, Erste Abteilung Originale~Reihe A 245: 171-183. Stipkovits, L., Z. Varga, G. Czifra, and M. Dobos Kovacs. 1986. Occurrence of mycoplasmas in geese affected with inflammation of the cloaca and phallus. Avian Pathology 15: 289-299. Taylor, R W. 1982. Seasonal aspects of the reproductive biology of the gopher tortoise, Gopherus polyphemus. Ph. D. Dissertation, University of Florida, Gainesville. 83 pp. TrampeL D. W., and O. J. Fletcher. 1981. Light microscopic, scanning electron microscopy, and histomorphometric evaluation of Mycoplasma gallisepticum induced airsacculitis in chickens. American Journal of Veterinary Research 42: 1281-1289. Trichard, C. J., P. A. Basson, J. J. van der Lugt, and E. P. Jacobsz. 1989. An outbreak of contagious bovine pleuropneumonia in the Owambo Mangetti area of South West Africa/Namibia: microbiological, immunofluorescent, pathological, and serological findings. Onderstepoort Journal of Veterinary Research 56: 277-284. Tully, J. G, D. L. Rose, J. B. Baseman, S. F. Dallo, A. L. LazzelL and C. P. Davis. 1995. Mixed Mycoplasma pneumoniae and Mycoplasma genitalium in a synovial fluid isolate. Journal of Clinical Microbiology 33: 1851-1855.

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108 U. S. Fish and Wildlife Service. 1990. Endangered and threatened wildlife and plants; determination of threatened status for the Mojave population of the desert tortoise. Federal Register 55: 12178-12190. Van Kuppeveld, F. J. M, J. T. M. Van Der Logt, A. F. Angulo, M. J. Van Zoest, W. G. V. Quint, H. G. M. Niesters, J. M. D. Galama, and W. J. G. Melchers. 1992. Genusand species-specific identification of mycoplasmas by 16s r RNA amplification. Applied Environmental Microbiology 58: 2602-2615. Washburn, L. R, and E. Weaver. 1996. Protective effect of active and passive immunization of rats against two surface antigens of mycoplasma arthritidis. IOM Letters 4: 31. Westhouse, R. A., E. R Jacobson, R. K. Harris, K. R. Winter, and B. L. Homer. 1996. Respiratory and pharyngo-esophageal iridovirus infection in a gopher tortoise (Gopherus polyphemus). Journal of Wildlife Diseases 32: 6812-686. Whittlestone, P. 1976. Effect of climatic conditions on enzootic pneumonia of pigs. International Journal of Biometeorology 20: 42-48. Wilson, D. S. 1991. Estimates of survival for juvenile gopher tortoises, Gopherus polyphemus. Journal of Herpetology 25: 376-379. Witz, B. W., D. S. Wilson, and M. D. Palmer. 1991. Distribution of Gopherus polyphemus and its vertebrate symbionts in three burrow categories. American Midland Naturalist 126: 152-158. Woldehiwet, Z., B. Mamache, and T. G. Rowan. 1990. Effects of age, environmental temperature, and relative humidity on the colonization of the nose and trachea of calves by Mycoplasma sp. British Veterinary Journal 146: 419-424. Wood, D. A. 1996. Florida's endangered species, threatened species, and species of special concern. Official lists. Florida Game and Fresh Water Fish Commission, Tallahassee, FL. 14 pp. Woodruff R E. 1982. Arthropods of gopher tortoise burrows. Pp. 25-48 in R. Franz and R J. Bryant (eds. ). The gopher tortoise and its sandhill habitat. Proceedings of the 3rd Annual Meeting Gopher Tortoise Council. Florida State Museum, Gainesville, FL. Wright, S. 1982. The distribution and population biology of the gopher tortoise (Gopherus polyphemus) in South Carolina. M. S. Thesis. Clemson University, Clemson, SC.

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109 Yoder, H. W., Jr. 1991. Mycoplasma gallisepticum infection, pp. 198-212 in: Calnek, B. W., H. J. Barnes, C. W. Beard, W. M. Reid, H. W. Yoder, Jr. (eds ). Diseases of Poultry. Iowa State University Press, Ames, IA. Yoder, H. W., Jr., and M. S. Hofstad. 1964. Characterization of avian mycoplasma. Avian Diseases 8: 481-512.

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BIOGRAPHICAL SKETCH GRACE SHERYL McLAUGHLIN I spent most of my life on the West Coast of North America, and attended Humboldt State University, Areata, California, receiving a Bachelor of Arts degree in Zoology, in spite of spending all my time in the Wildlife Department. I then ran an organic farm for several years before moving to Ames, Iowa, to attend Iowa State University. I studied gopher tortoises on Sanibel Island, Florida, en route to a Master of Science degree in Animal Ecology. Before defending my thesis in November 1990, 1 began my doctoral studies at University of Florida. I originally studied parasites of bobcats and Florida panthers, particularly their hookworms. However, I accepted a job with the gopher tortoise upper respiratory disease project, and switched dissertation research projects. I hope to move west and/or north soon after completion of my work on the project. 110

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Donald J. Forrester, Chair Professor of Wildlife Ecology and Conservation I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Mary B. Brown, Cochair Associate Professor of Veterinary Medicine I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Melvin E. Sunquist 0 Associate Professor of Wildlife Ecology and Conservation I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Wiley M. Kj(§hens Associate Professor of Wildlife Ecology and Conservation I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. <3$5Uggg fc^ttftfefr Elliott R. Jacobson Professor of Veterinary Medicine

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I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. ^ aid Paul A. Klein Professor of Pathology, Immunology and Laboratory Medicine This dissertation was submitted to the Graduate Faculty of the College of Agriculture and to the Graduate School and was accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy. /") May 1 997 ^ Dean, College of Agriculture Dean, Graduate School


78
of the 1995 samples. Additionally, that modification reduced the necessary blood volume
from approximately 400 pi to 150 pi, reducing stress to the hatchlings. Samples collected
in 1995 and 1996 from the corresponding female were run on the same plate as samples
from their eggs and/or hatchlings.
Results
Clutch Sizes. Fertility and Hatching Rates
Twenty-six clutches, 13 each year, were collected. Arginine vasotocin was used to
induce opposition of 20 clutches and the remaining six clutches (five in 1995 and one in
1996) were laid in the pens. In 1995, 13 tortoises developed 115 eggs (mean = 8.8), and
103 eggs were recovered (Table 6-1). Five females laid entire clutches in the pens, four of
which were removed within 1 wk of laying. One female laid nine eggs in an inaccessible
area of her pen. Two females that laid incomplete clutches on induction laid the remaining
eggs (w 3) in their pens. Twenty-five eggs were removed for culture, PCR, and ELISA
samples. Of those, five were fertile and removed while Pable, four had died, and five
were infertile. I could not determine fertility on the remaining 11.
Overall clutch sizes (Table 6-1) were larger in 1996 (mean = 11.1) than in 1995
(mean = 8.8, 2-tail /-test = 2.78, 17 df P = 0.013), as were those of seronegative females
(11.7 vs 9.8, 2-tail /-test = 2.45, 9 df, P = 0.037). In 1996, clutch sizes of seronegative
females (mean = 11.7) were larger than those of seropositive females (mean = 10.3, 2-tail
/-test = 2.30, 11 df, P = 0.042). Thirteen females developed 144 eggs, 130 of which were
recovered. One female laid 11 eggs that were incompletely calcified and broke during the


29
The following criteria were utilized for grading lesions:
Normal (score = 0): Occasional small subepithelial lymphoid aggregates; rare
heterophils in the lamina propria. No changes in mucosal or glandular epithelium; no
edema.
Mild (1): Multifocal small subepithelial lymphoid aggregates; multifocally, small
numbers of heterophils, lymphocytes, and plasma cells in the lamina propria; mild edema in
lamina propria; minimal changes in mucosal epithelium.
Moderate (2-3): 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; proliferation and disorganization of the basal epithelium.
Severe (4-5): Focally extensive to diffuse band of lymphocytes and plasma cells
subjacent to and obscuring the overlying mucosal epithelium; large numbers of heterophils
in lamina propria and infiltrating 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; occasional squamous metaplasia.
Statistical Analyses
Statistical analyses were performed using SAS (SAS Institute, 1988) or SigmaStat
for Windows, Version 1.0 (Fox et al. 1994). Because the analyses varied for each
experiment, the specific methods will be addressed in the appropriate chapters.


DEDICATION
This dissertation, the years that went into it, and the drive for knowledge
that underpinned it, are a result of, and dedicated to, my parents:
Grace Slater Gibson Billie McLaughlin
1928-1978
and
Robert Douglas McLaughlin
1925-1994
Thank you.


10
or very cold winters. Human impacts on tortoises and their habitat, whether through
disruption of normal behavior patterns, degradation of habitat through agriculture,
silviculture, mining or development operations, or pollution, may cause sufficient
physiological stress to trigger proliferation of the mycoplasma and recurrence of signs.
Capturing and transporting of tortoises during relocation, restocking and repatriation
efforts also may be significant sources of stress.
Mycoplasmal Respiratory Diseases in Domestic Animals and Humans
Mycoplasmas cause respiratory disease in other taxa, including rodents, poultry,
swine, ruminants, and humans. All have similar pathological effects, which are described
below, and many are exacerbated by concurrent infections or environmental stress.
Murine respiratory disease caused by Mycoplasma pulmonis has caused problems
in laboratory settings for more than 70 yr (Lindsey et al. 1971). It has necessitated
complex and expensive control measures, including various types of isolation mechanisms
and cesarean delivery of mice and rat pups to reduce the prevalence of M. pulmonis
infections in colonies (Cassell et al. 1984, Davis et al. 1987). Interactions among
environment, host, and strain factors influence the impacts at individual and population
levels (Simecka et al. 1992). Histologically, lesions are characterized by lymphoid
hyperplasia and chronic inflammation (Cassell et al. 1985).
Poultry respiratory diseases can be caused by viruses, mycoplasmas, and other
bacteria, singly or in combination. Without concurrent viral or other bacterial infections,
disease can be mild or subclinical (Kerr and Olson 1967). Environmental factors,
particularly dust and ammonia levels, as well as strain differences, affect the severity of


CHAPTER 1
INTRODUCTION
Gopher Tortoise Natural History
Gopher tortoises, Gopherus polyphemus, are found in the southeastern United
States, on the coastal plain from southern South Carolina south through Georgia and
throughout Florida, and west through southern Alabama, Mississippi, and Louisiana. The
major population concentrations are in Florida and southern Alabama and Georgia, with
only remnant populations in South Carolina, Mississippi and Louisiana (Auffenberg and
Franz 1982, Diemer 1992a). Populations are concentrated in areas with deep sandy soils
suitable for digging. Vegetation associations in which tortoises are found include longleaf
pine-xerophytic oak woodlands, palmetto scrub, sand pine scrub, oak scrub, beach scrub,
coastal strands, pine flatwoods, dry prairies, native pasture, and savanna, as well as ruderal
habitats (Landers and Speake 1980, Lohoefener and Lohmeier 1981, McRae et al. 1981,
Campbell and Christman 1982, Diemer 1986, Breininger et al. 1988).
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 (Eisenberg 1983).
Gopher tortoises live in loose colonies, with considerable movement of tortoises among
groups over the years (Diemer 1992b). Colonies may be defined more by the availability
of suitable soils for digging burrows, or the distribution of food resources, than by social
1


11
outbreaks of mycoplasmosis. Other stress factors, such as crowding and cold weather,
also influence morbidity and mortality (Jordan 1972). Mycoplasma gallisepticum causes
chronic respiratory disease in chickens and sinusitis in turkeys, and affects ring-necked
pheasants (Phasianus colchicus), chukar (Alectoris chukar), northern bobwhite (Colinus
virginianus), common peafowl (Pavo cristatus) (Yoder 1991) and Japanese quail
(Coturnix japnica) (do Nascimento and do Nascimento 1986). Lower respiratory tract
complications are seen rarely in turkeys (Simecka et al. 1992). Histologically, hypertrophy
and hyperplasia of respiratory epithelia, including mucous cells, are seen, as is lymphoid
infiltration of the submucosa. Heterophilic exudate is seen in the airways (Nunoya et al.
1987, Trampel and Fletcher 1981). Mycoplasma meleagridis infection is seen primarily in
chicks and poults up to 10 wks of age, and is sexually transmitted. Turkeys, Japanese
quail, and peafowl develop air sacculitis, sometimes with accompanying tracheitis and
pneumonia, but not sinusitis. Histologically, the lesions are characterized by hypertrophy
and hyperplasia of the air sac epithelia, with edema and lymphoid infiltration (Stipkovits
1979). Young chickens and turkeys also are susceptible to M. synoviae, usually in
conjunction with Newcastle disease virus or infectious bronchitis (Hopkins and Yoder
1984, Springer et al. 1974). As with M. gallisepticum, turkeys develop sinusitis
(Stipkovits 1979). All three bacteria can be transmitted vertically (i.e., via the egg)
(Simecka et al. 1992).
Swine develop mild pneumonia when infected by M. hyopneumoniae, and although
mortality is virtually nil, the disease is chronic, causing slow growth and reduced weight
gains, decreasing profitability (Oboegbulem 1981, Jericho 1986). Although signs can
disappear, recurrences will occur with weather changes, viral infections, and other


77
frozen for later analyses. For culture, 100 |il samples of yolk and albumin, and small
pieces of membrane were added to 900 pi of SP4 broth. After 48 hr incubation at 30C,
500 pi of culture were removed for PCR analysis. All CAF samples obtained at pipping
were treated in the same manner. In 1996, one or two pooled samples consisting of 100
pi of CAF from each of 3 6 eggs, mixed well, were made for each clutch and processed
as described.
ELISA Procedures
Based on preliminary experiments using several methods of antibody extraction,
the supernatant resulting from mixing 1 ml of yolk with 1 ml PBS-AZ provided the most
efficient fraction for detection of antibodies (G. S. McLaughlin and I. M. Schumacher,
unpub. data). Blood samples for ELISA were collected by cardiocentesis from most
hatchlings at approximately 2-4 wk of age. In 1995, the goal was to obtain a blood
sample from each hatchling. In 1996, sampling effort was reduced to a maximum of five
samples per clutch due to the low coefficients of variation for the 1995 samples.
In 1995, hatchling ELISA samples were run as previously described except that
three dilutions from the range 1:1 to 1:8 were run in duplicate, depending on the sample
volume obtained. Sample dilutions were based on previous experiments with plasma from
desert tortoise hatchlings (I. M. Schumacher, unpub. data). All hatchling or egg samples
were run on the same plate as 1994 and 1995 samples from the corresponding female and
the presumed sire, if possible. If samples were split between two plates, plates were run
on the same day with the same reagents and positive and negative controls. In 1996,
samples were run in triplicate at a dilution of 1:2, as that was the dilution used for analysis


44
toxic compounds or induce the production of compounds by the tortoises that cause
damage to the liver. Desert tortoises with URTD also show changes in liver tissue (B. L.
Homer, pers. comm.), and some exhibit altered nitrogen metabolism (B. Henen, pers.
comm.). The altered nitrogen metabolism may be due to behavioral changes leading to
decreased foraging rates or efficiency, or it may be due to direct effects of the
Mycoplasma infection, but the mechanism is not yet understood. Although only one of
nine experimentally infected captive gopher tortoises significantly decreased its intake of
vegetables (G. S. McLaughlin, unpub. data), and that decrease was temporary, wild
tortoises may alter their behavior patterns to a greater degree. Alternatively, secondary
infections by other bacteria may be the proximate cause of liver damage. However, no
evidence of primary or secondary bacterial infection (i.e., necrosis) was seen histologically.
By the ELISA, none of eight clinically healthy tortoises and 11 of 15 diseased
tortoises had antibodies against M. agassizii, indicating previous exposure. Tortoises with
negative ELISA results may have been in the early stages of the disease, when increased
antibody levels had not occurred or were not detectable. The four diseased, seronegative,
tortoises may have been infected with another agent. Westhouse et al. (1996) implicated
an iridovirus as the cause of pneumonia, tracheitis, pharyngitis, and esophagitis in a gopher
tortoise from Sanibel Island. The virus was readily detectable on both light and electron
microscopy. It is not known if attempts were made to culture mycoplasma, or if an
ELISA was run on a plasma sample. No indications of viral infections were seen in the
tortoises examined for the current study.
Investigators recently have found seropositive gopher tortoises in Georgia (B.
Raphael, pers. comm.), seropositive, clinically ill, and/or PCR positive tortoises at a site in


7
Clinical signs of URTD in gopher and desert tortoises include serous, mucoid, or
purulent discharge from the nares, excessive tearing to purulent ocular discharge,
conjunctivitis, and edema of the eyelids and ocular glands (Jacobson et al. 1991, G. S.
McLaughlin personal observations). Individual infected tortoises vary in the suite of signs
they have, and the severity can vary from day to day. Nares may become occluded with
caseous exudate, preventing externally visible nasal discharge. Lymphocytic infiltration of
the comeas, while rare, may decrease an animals ability to forage or avoid predators.
Tortoises may become lethargic and anorectic, leading to dehydration, emaciation, and
eventual death from cachexia. Lethargy, nonresponsiveness to stimuli, and altered
behavior pattemssuch as basking at lower temperatures than normalmay render a
tortoise more susceptible to predation. Moribund animals often develop petechial to
ecchymotic hemorrhages under the scutes, especially visible on the plastron (G. S.
McLaughlin pers. obs.), which may be due to septicemia caused by secondary infection
with opportunistic bacteria.
Several agents were hypothesized to cause respiratory tract disease in tortoises,
including viruses, Mycoplasma sp. (Lawrence and Needham 1985), and Pasteurella
testudinis (Snipes and Biberstein 1982). However, experimental infections to determine
the etiologic agent were not conducted. Beginning in 1989, efforts were undertaken to
determine the etiology of URTD. Clinically ill and healthy desert tortoises from California
were examined and samples collected, either from live animals or at necropsy.
Hematologic and serum biochemical evaluations, liver vitamin and metal determinations,
and pathologic and microbial investigations were conducted (Jacobson et al. 1991).


68
Transmission Probabilities
Analysis of the data was complicated by the changing exposure status of the
tortoises due to the housing problems discussed above. Therefore, analyses were carried
out on three data sets. The first consisted of data from the first 10 mo (August 1994 -
June 1995) of the study, the second from the last 14 mo (June 1995 July 1996), and the
third of the cumulative data. The first data set (Table 5-2) consisted of the original 15
pairs. The second set (Table 5-3) did not include those pairs in which transmission had
already occurred, but only the control pairs and those whose exposure status had changed,
leaving 11 observations. The third set (Table 5-4), with 22 observations, included some
tortoises twice, as the observations of interest were the exposure events themselves, and
not the pairs or individual tortoises. Although sample sizes were very small, and some
cells had no observations, chi-squared tests of differences in proportions were run on the
three data sets. The final set was collapsed on the three predictor variables (ELISA,
clinical, and culture/PCR status; Table 5-5) to determine which had the most influence on
transmission probability.
Four exposure status classes were established based on ELISA classification,
clinical signs, and culture or PCR status. Class assignment was based on a tortoises
status at any point during the time period of interest. Therefore, a tortoises status could
change between the first and last parts of the study. Each criteria was recorded as positive
or negative, and transmission was recorded as yes or no. Transmission was classified very
conservatively, with any tortoise exhibiting even mild, transient, clinical signs not
obviously related to mechanical irritation (e.g., plant material in the eye or nose) or
environmental conditions (e.g., being held overnight in a box in which the tortoise had


43
changes in the upper respiratory tract of the gopher tortoises were similar to inflammatory and
dysplastic changes reported for desert tortoises with URTD (Jacobson et al. 1991, Brown et
al. 1994, Jacobson et al. 1995). However, inflammationa and epithelial proliferation around
the glottis, tracheitis, and proliferative pneumonia have not been seen in desert tortoises.
Additionally, two diseased tortoises had proliferation of the colonic mucosal epithelium.
Those changes have not been seen in desert tortoises with URTD.
By electron microscopy, organisms consistent with Mycoplasma were
demonstrated on the nasal mucosal surfaces of two tortoises. Other than bacteria, no
infectious agents were demonstrated in or on nasal cavity mucosa by electron microscopy.
Eleven diseased gopher tortoises were PCR positive and M. agassizii was cultured from
the upper respiratory tract of 11 diseased tortoises examined. These results support the
hypothesis that M. agassizii is a cause of URTD in gopher tortoises in Florida.
The greater number of species and increased proportion of Gram-negative bacteria
isolated from diseased tortoises could indicate that conditions in the upper respiratory
tract of diseased tortoises are more favorable for the growth of those species, or that
tortoises infected with M. agassizii are more susceptibile to opportunistic pathogens. The
positive culture and / or PCR results from nasal cavity samples (obtained at necropsy) of
four clinically healthy tortoises with negative nasal passage flushes and swabs support the
hypothesis that tortoises can harbor the organism without showing clinical signs or
shedding bacteria. Such animals may recrudesce under stressful conditions, begin
shedding bacteria, and become infective to other tortoises.
Nine gopher tortoises exhibited pathologic changes in the livers, although the
significance of these changes is not understood currently. The mycoplasma may release


76
Egg Collection and Incubation
Egg development was monitored by radiography (6 mas, 62-76 kv) beginning in
April of 1995 and 1996. When eggshells were judged to be calcified adequately,
oviposition was induced by intravenous injection of arginine vasotocin (Sigma) at
approximately 10-12 picograms/kg. Tortoises were monitored until clutch deposition was
complete or for a minimum of 4 hr. Eggs were collected as they were laid and placed in a
sanitized container (washed with 1:10 bleach solution and air-dried upside down) partially
filled with sterilized vermiculite moistened with an equal weight of sterilized water.
Female identification number and letter indicating order of deposition were written on
each egg with a graphite pencil swabbed with 70% ethanol. After approximately half the
clutch was laid, cloacal swabs for mycoplasma culture were taken, streaked onto SP4
agar, placed in SP4 broth, and incubated as described previously.
In 1995, eggs were incubated at 29C until hatching. In 1996, eggs were
incubated at 27, 29 or 31C. Approximately 1 wk before hatching each egg was cleansed
of adhering vermiculite with clean gauze and placed into a sanitized plastic container.
After hatching, resorption of yolk, and closing of the umbilicus, hatchlings were placed in
containers with clean sand. Hatchlings were maintained separately from adults, at ambient
temperature and light cycles, and fed natural foods supplemented with commercially
available vegetables until release into outdoor pens, where they were fed natural foods.
Culture and PCR Procedures
Eggs were taken at various times during incubation for mycoplasma culture and
antibody detection. At pipping, chorioallantoic-amniotic fluid (CAF) was collected and


53
Culture and PCR Results
The sentinel tortoise that became ill also had positive culture and PCR results. All
inoculated tortoises had at least two positive culture results, and 17 of 18 had positive
cultures from samples collected at necropsy. Mycoplasmal DNA was detected by PCR
analysis from each inoculated tortoise, including those that did not exhibit clinical signs of
URTD when sampled. By RFLP analysis, all isolates corresponded to M. agassizii.
ELISA Results
There was no anti-M. agassizii antibody response by the control tortoises to sham
inoculation, although one sentinel had seroconverted by 4 wk PI. All inoculated tortoises
seroconverted or had significantly increased ELISA values. ELISA values were greater
for challenge than naive tortoises at each time point (Table 4-2, Figure 4-3). A significant
increase in ELISA values of challenged tortoises was observed by 4 wk PI (mean ratio of
samples to negative control of 6.3 vs. 3.3 at 0 wk, P < 0.05), and seroconversion of naive
tortoises was observed by 6 wk PI (ratio of 3.9 vs. 1.5 at 0 wk, P < 0.05). The ratio of
the challenge to the naive ELISAs increased at 2 and 4 wk PI, then declined.
Table 4-2. Least-squares mean ELISA values for naive and challenge tortoises. P < 0.05
indicates a significant difference between groups.
Time
Naive
Challenge
P
0 weeks pi
0.2049
0.4707
0.0437
2 weeks pi
0.2235
0.5925
0.0126
4 weeks pi
0.2439
0.8848
0.0001
6 weeks pi
0.5464
1.5254
0.0001
8 weeks pi
0.8314
1.8219
0.0001
12 weeks pi
1.4062
1.9839
0.0001
Necropsy
1.6796
2.2889
0.0001


27
a pen. Animals were captured by hand or using wire cage-type traps (Tomahawk Live
Trap Company, Tomahawk, WI) that were covered with brown paper to protect the
animals from the weather. Traps were cleaned, sprayed with bleach solution, and allowed
to air dry following each use. 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, WI) for transport and holding. Containers were
bleached, scrubbed, and washed in an automatic cage washer before reuse.
D C
A B
Figure 2-1. Layout of tortoise pens at the University of Florida Animal Resource Farm.
The outer fence was chain link, and the inner fence and pen dividers were corrugated sheet
metal on a wooden frame.
Necropsy Procedures
All diseased and selected healthy tortoises were euthanatized with a combination of
drugs. Ketamine was administered intramuscularly at 60 80 mg/kg followed by a


107
Schumacher, I. M, M. B. Brown, E. R. Jacobson, B. R. Collins, and P. A. Klein. 1993.
Detection of antibodies to a pathogenic mycoplasma in desert tortoises (Gopherus
agassizii) with upper respiratory tract disease. Journal of Clinical Microbiology, 31:
1454-1460.
Simecka, J. W., J. K. Davis, M. K. Davidson, S. E. Ross, C. T. K. H. Stadtlander, and G.
H. Cassell. 1992. Mycoplasmas which infect animals. Pp. 391-415 in: Maniloff, J.,
R. N. McElhaney, L. R. Finch, J. B. Baseman, eds. Mycoplasmas: molecular biology
and pathogenesis. American Society for Microbiology, Washington, D. C.
Snipes, K. P., and E. L. Biberstein. 1982. Pasteurella testudinis sp. nov.: a parasite of
desert tortoises. International Journal of Systematic Bacteriology 32: 201-210.
Springer, W. T., C. Luskus, and S. S. Pourciau. 1974. Infectious bronchitis and mixed
infections of Mycoplasma synoviae and Escherichia coli in gnotobiotic chickens. I.
Synergistic role in the airsacculitis syndrome. Infection and Immunity 10: 578-589.
Stalheim, O. H. 1983. Mycoplasmal respiratory diseases of ruminants: a review and
update. Journal of the American Veterinary Medical Association 182: 403-406.
Stipkovits, L. 1979. The pathogenicity of avian mycoplasmas. Zentralblatt fur
Bakteriologie, Para sit enkunde Infektionskrankheiten und Hyggiene, Erste Abteilung
OriginaleReihe A 245: 171-183.
Stipkovits, L., Z. Varga, G. Czifra, and M. Dobos Kovacs. 1986. Occurrence of
mycoplasmas in geese affected with inflammation of the cloaca and phallus. Avian
Pathology 15: 289-299.
Taylor, R. W. 1982. Seasonal aspects of the reproductive biology of the gopher tortoise,
Gopherus polyphemus. Ph. D. Dissertation, University of Florida, Gainesville. 83 pp.
Trampel, D. W., and O. J. Fletcher. 1981. Light microscopic, scanning electron
microscopy, and histomorphometric evaluation of Mycoplasma gallisepticum induced
airsacculitis in chickens. American Journal of Veterinary Research 42: 1281-1289.
Trichard, C. J., P. A. Basson, J. J. van der Lugt, and E. P. Jacobsz. 1989. An outbreak of
contagious bovine pleuropneumonia in the Owambo Mangetti area of South West
Affica/Namibia: microbiological, immunofluorescent, pathological, and serological
findings. Onderstepoort Journal of Veterinary Research 56: 277-284.
Tully, J. G., D. L. Rose, J. B. Baseman, S. F. Dallo, A. L. Lazzell, and C. P. Davis. 1995.
Mixed Mycoplasma pneumoniae and Mycoplasma genitalium in a synovial fluid
isolate. Journal of Clinical Microbiology 33: 1851-1855.


98
Bradbury, J. M, A. VuiUaume, J. P. Dupiellet, M. Forrest, J. L. Bind, G. Gaillard-Perrin.
1987. Isolation o Mycoplasma cloacale from a number of different avian hosts in
Great Britain and France. Avian Pathology 16: 183-186.
Breininger, D. R., P. A. Schmalzer, D. A. Rydene and C. R. Hinkle. 1988. Burrow and
habitat relationships of the gopher tortoise in coastal scrub and slash pine flatwoods
on Merritt Island, Florida. Final report. Florida Game Fresh Water Fish Commission,
Tallahassee, FL. 238 pp.
Brown, D. R., T. L. Clippinger, K. E. Helmick, I. M. Schumacher, R. A. Bennett, C. M.
Johnson, K. A. Vliet, E. R. Jacobson, and M. B. Brown. 1996a. Mycoplasma
isolation during a fatal epizootic of captive alligators (.Alligator mississippiensis) in
Florida. IOM Letters 4: 42-43.
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. International Journal of Systematic Bacteriology 45: 348-350.
Brown, D. R., G. S. McLaughlin, I. M. Schumacher, M. B. Brown, E. R. Jacobson, and P.
A. Klein. 1996b. Dose response study of Mycoplasma agassizii infection in gopher
tortoises. Desert Tortoise Council 1996 Annual Meeting and Symposium Abstracts,
10.
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. Infection and Immunity 62: 4580-4586.
Bryson, D. G. 1985. Calf pneumonia. Veterinary Clinics of North America Food Animal
Practice 1: 237-257.
Campbell, H. W., and S. P. Christman. 1982. The herpetological components of Florida
sandhill and sand pine scrub associations. Pp. 163-171 in N. J. Scott, Jr. (ed.).
Herpetological Communities. U. S. Fish Wildlife Service, Wildlife Research Report
13.
Carmichael, L. E., T. D. St. George, N. D. Sullivan, and N. Horsfall. 1972. Isolation,
propagation, and characterization studies of an ovine mycoplasma responsible for
proliferative interstitial pneumonia. Cornell Veterinarian 62: 654-679.
Cassell, G. H., W. A. Clyde, Jr., and J. K. Davis. 1985. Mycoplasmal respiratory
infections. Pp. 65-106 in S. Razin and M. F. Barile (eds.). The mycoplasmas, vol. 4.
Mycoplasma pathogenicity. Academic Press, New York.


32
identification systems API 20E for enteric organisms and API NFT for non-enterics
(BioMerieux Vitek, Hazelwood, MO, USA). Isolates of organisms consistent with
Pasteurella were identified to species according to biochemical profiles fisted for P.
testudinis (Snipes and Biberstein 1982).
Electron Microscopy
Selected specimens were submitted to H. P. Adams of New Mexico State
University, Las Cruces, for scanning and transmission electron microscopic preparation
and evaluation.
The left half of the bisected head of one healthy tortoise was prepared for
ultrastructural evaluations. The nasal cavity was instilled with 2.5% glutaraldehyde in 0.1
M phosphate buffer, then dissected out in its entirety, and selected areas were sampled.
The tissues were dehydrated in an ascending series of ethanols and transferred to
hexamethyldifisilazane for the final drying. The samples were sputter coated with gold and
viewed by scanning electron microscopy (SEM).
Samples from 11 diseased tortoises were collected for transmission electron
microscopy (TEM). The nasal cavity tissue was removed from the underlying
cartilaginous tissues and separated into anterior dorsal, anterior ventral, posterior dorsal
and posterior ventral quadrants. Each quadrant was cut into 1 mm cubes and placed in
2.5% glutaraldehyde, and post-fixed in osmium tetroxide. Specimens were prepared for
TEM by embedding in epon-araldite and sectioning with an ultramicrotome. Thick
sections were stained with toluidine blue and examined by fight microscopy. Ultrathin
sections were placed on copper grids, stained with uranyl acetate and lead citrate, and
examined with a Hitachi H7000 transmission electron microscope.


CHAPTER 2
METHODS
Tortoises. Intake Procedures. Clinical Assessments and Sampling Methods
For the natural infection studies, tortoises were obtained from various locations in
the state of Florida under Florida Game and Fresh Water Fish Commission permits
number WX93227 issued to Elliott R. Jacobson and number WX94037 issued to Mary B.
Brown. Tortoises were processed the day of arrival at the University of Florida (UF),
Gainesville. For the natural and experimental infection studies, gopher tortoises were
transferred from a development site in central Florida to UF in April, July, and August
1994 and April 1995, and processed the day following arrival. Tortoises were examined
for clinical signs of URTD: nasal and ocular discharge, palpebral edema, and
conjunctivitis. The signs were graded individually on a scale of 0-3, none to severe. Total
clinical sign score was calculated as the nasal discharge score plus the mean of three
ocular sign scores (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. Straight line
carapace length, thickness, and width were measured to the nearest cm with forestry
calipers.
21


38
Figure 3-4. Photomicrograph of the nasal cavity tissues of a gopher tortoise with upper
respiratory tract disease. The changes were classified as severe, with
aggregates of lymphoid cells in the submucosa, proliferation of the basal cells,
and dysplasia of the mucosal epithelium. Hematoxylin and eosin staining,
320x. Photograph by E. R. Jacobson.


18
Chronic Manifestations of Mycoplasmal Infections
Many mycoplasmal diseases are characterized by an overaggressive or
inappropriate immune response by the host, eventually leading to autoimmune damage to
the affected sites, whether respiratory or urogenital tract, joints, heart, skin, or other organ
systems (Krause and Taylor-Robinson 1992, Simecka et al. 1992, Cole 1996). Infected
hosts may be more susceptible to secondary infections with other bacteria or viruses. In
wildlife species, such complications may reduce the fitness of the animals by altering
behavior, leading to decreased foraging efficiency, increased susceptibility to predators, or
diminished mate seeking behavior. Energy that normally would be allocated to
reproduction may be needed to repair or compensate for damage to multiple organ
systems. Therefore, even if mycoplasmal infections do not cause mortality directly, they
can affect individual and population viability.
Project Overview and Specific Objectives
Due to the listing by the Florida Game and Fresh Water Fish Commission of the
gopher tortoise as a species of special concern, and the subsequent permitting of over 450
relocations involving more than 8000 tortoises, particular attention has been focused on
the dynamics and persistence of both natural and relocated populations. Understanding
the effects of URTD on individuals and populations is essential for proper management of
remaining populations; therefore, a study was begun in 1993 on the etiology, pathology,
and diagnosis of URTD in gopher tortoises. The original objectives were 1) to describe
the pathology of natural infections and identify possible etiologic agents, 2) to perform


UPPER RESPIRATORY TRACT DISEASE IN GOPHER TORTOISES,
GOPHERUS POLYPHEMUS:
PATHOLOGY, IMMUNE RESPONSES, TRANSMISSION,
AND IMPLICATIONS FOR CONSERVATION AND MANAGEMENT
BY
GRACE SHERYL MCLAUGHLIN
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1997


58
on this study, tortoises testing positive for antibodies against M agassizii cannot be
considered good candidates for release in repatriation, restocking, or relocation efforts. It
may be acceptable to release seropositive tortoises in areas that already have seropositive
animlas, as long as the overall prevalence is not increased significantly. Seropositive
tortoises may provide desirable genetic material (see Chapter 6), and would be valuable as
research subjects to study interactions with other disease agents and the effects of
repeated exposure on metabolism, autoimmune responses, reproduction, and survival.


40
ELISA and PCR Results
ELISA and PCR results for both healthy and diseased tortoises are presented in
Table 3-1. All healthy gopher tortoises were seronegative for antibody specific against M.
agassizii. Twelve diseased tortoises were seropositive, two were suspect, and one was
seronegative. All healthy tortoises were PCR negative while 11 diseased tortoises were
PCR-positive for Mycoplasma in nasal aspirates. Four clinically healthy tortoises that had
negative culture and PCR results from nasal passage flush and swab samples had positive
cultures and / or PCR results for samples from the nasal cavities.
Microbial Isolation Results
The results of Mycoplasma and aerobic microbial cultures of the upper respiratory
tract (URT) are presented in Tables 3-1 and 3-2. Mycoplasma was not cultured from the
URT of any healthy tortoise. Mycoplasma was cultured from the URT of 11 tortoises
with URTD. The aerobic microbial isolates of healthy tortoises consisted primarily of
members of the genera Staphylococcus, Streptococcus, and Corynebacterium a few
Gram-negative rods were isolated. A greater number of Gram-negative species were
isolated from the nasal cavities of tortoises with URTD, and those isolates made up a
greater proportion of the isolates. Pasteurella testudinis was isolated from five diseased
tortoises; in two it represented the major aerobic isolate.
Discussion
Gopher tortoises with clinical signs of URTD and evidence of exposure to
Mycoplasma agassizii were obtained from multiple sites in Florida. The light microscopic


2
interactions (Campbell and Christman 1982). Gophers are the most fossorial of the four
North American species of tortoises, digging burrows that may extend 5 meters down
from the surface and 15 meters in length (Hansen 1963, Diemer 1986). The burrows
provide a microclimatically stable environment for not only the tortoises, but also for
numerous commensals. Approximately 60 vertebrate speciesfrom snakes to birdsand
over 300 invertebratesincluding spiders, crickets, and beetleshave been found in
tortoise burrows or observed using them as permanent homes or refuges from heat, cold,
fire, and predators (Jackson and Milstrey 1989, Lips 1991, Witz et al. 1991). Some
invertebrate species are obligate commensals, and occur only in active tortoise burrows,
where they feed on tortoise feces and other invertebrates and, in turn, are eaten by other
commensals, such as gopher frogs (Rana areolata) and Florida mice (Podomys
floridanus) (Woodruff 1982). Several species that exclusively or frequently use tortoise
burrows have legal protection in Florida and other parts of their ranges. These include
scarab beetles (F. Scarabaeidae), indigo (Drymarchon coris couperi) and pine (Pituophis
melanoleucus) snakes, gopher frogs, mole skinks (Eumeces egregius), burrowing owls
(Athene cunicularia floridana), and Florida mice (Cox et al. 1987).
The soil disturbance resulting from the digging of the burrows allows deeper
access of air and water into the soil profile, as well as providing bare mineral soil patches
on the surface. When a burrow is abandoned the soil mound, or apron, in front of the
entrance no longer undergoes continual disturbance, allowing certain plants to colonize
the area. The composition of the plant assemblage on the mound may differ from that in
the surrounding undisturbed area, providing a mosaic of small patches in the habitat
(Breininger et al. 1988, McLaughlin 1990). Tortoises consume a wide variety of grasses


104
Ley, D. H., J. E. Berkhoff, K. Joyner, L. Powers, and S. Levisohn. 1996. Molecular
epidemiology investigations of Mycoplasma gallisepticum conjunctivitis in songbirds
by random amplified polymorphic DNA (RAPD) analysis. IOM Letters 4: 53-54.
Ley, D. H., J. E. Berkhoff, and J. M. McLaren. 1996. Mycoplasma gallisepticum
isolated from house finches (Carpodacus mexicanus) with conjunctivitis. Avian
Diseases 40: 480-483.
Lindsey, J. R., H. J. Baker, R. G. Overreach, G. H. Cassell, and C. E. Hunt. 1971.
Murine chronic respiratory tract disease: Significance as a research complication and
experimental production with Mycoplasma pulmonis. American Journal of Pathology
64: 675-708.
Linley, T. A., and H. R. Mushinsky. 1994. Organic composition and energy content of
eggs and hatchlings of the gopher tortoise. Pp. 113-128 in R. B. Bury and D. J.
Germano (eds.). Biology of North American tortoises. National Biological Survey,
Fish and Wildlife Research 13.
Lips, K. R. 1991. Vertebrates associated with tortoise (Gopherus polyphemus) burrows
in four habitats in south-central Florida. Journal of Herpetology 25: 477-481.
Lohoefener, R., and L. Lohmeier. 1981. Comparison of gopher tortoise (Gopherus
polyphemus) habitats in young slash pine and old longleaf pine areas of southern
Mississippi. Journal of Herpetology 15:239-242.
Luttrell, M. P., S. H. Kleven, and W. R. Davidson. 1991. An investigation of the
persistence of Mycoplasma gallisepticum in an eastern population of wild turkeys.
Journal of Wildlife Diseases 27: 74-80.
Luttrell, M. P., J. R. Fischer, D. E. Stallknecht, and S. H. Kleven. 1996. Field
investigation of Mycoplasma gallisepticum infections in house finches (Carpodacus
mexicanus) from Maryland and Georgia. Avian Diseases 40: 335-341.
Lyell, A., A. M. Gordon, H. M. Dick, and R. G. Sommerville. 1967. Mycoplasmas and
erythema multiforme. Lancet 2: 1116-1118.
Macdonald, L. A., and H. R. Mushinsky. 1988. Foraging ecology of the gopher tortoise,
Gopherus polyphemus, in a sandhill habitat. Herpetologica 44:345-353.
Mann, T. M. 1990. The status of Gopherus polyphemus in South Carolina. Pp. 88-130
in C. K. Dodd, Jr., R. E. Ashton, Jr., R. Franz, and E. Wester (eds.). Proceedings of
the 8th Annual Meeting Gopher Tortoise Council, Florida Museum of Natural
History, Gainesville, FL.
Mansel, J. K., E. C. Rosenow III, T. F. Smith, and J. W. Martin, Jr. 1989. Mycoplasma
pneumoniae pneumonia. Chest 95: 639-646.


CHAPTER 7
ENVIRONMENTAL TRANSMISSION OF MYCOPLASMA AGASSIZII
Introduction
Some mycoplasmas can remain viable for at least several days in the environment
(Chandiramani et al. 1966), and for many years under refrigeration or freezing (Yoder and
Hofstad 1964). One method of controlling mycoplasma infections in domestic stock is to
depopulate a farm or facility, spray buildings and equipment with disinfectants, wait an
appropriate amount of time, and then reintroduce uninfected stock (Anonymous 1989). In
order to determine appropriate time frames for restocking tortoises, knowing the length of
time Mycoplasma agassizii remains viable in the environment is critical. Unfortunately,
direct viability testing is extremely difficult and impractical, if not impossible. Many soil
bacteria and fungi in the tortoise burrow and surrounding environment grow very rapidly
in culture and quickly overtake any mycoplasma colonies that might be present. However,
the risk of environmental transmission of M. agassizii is an important parameter in the
decision making process. In order to address this question, I designed an experiment to
test the hypothesis that environmental transmission ofM agassizii would occur in
tortoises introduced to pens previously occupied by infected, clinically ill tortoises.
84


15
wild, free-ranging birds, and their potential impacts on the populations need to be further
investigated.
Several species of mycoplasmas, including M. cloacale (Bradbury et al. 1987,
Goldberg et al. 1995) and M. anatis (Ivanics et al. 1988, Poveda et al. 1990, Goldberg et
al. 1995), have been isolated from semi-domestic and wild ducks and other avian species
throughout the world. Stipkovits et al. (1986) reported isolation ofM cloacale from
geese with inflammation of the cloaca and phallus, but Goldberg et al. (1995) found no
association ofM cloacale with disease in wild mallards (Anas platyrhynchos), black
ducks (A. rubripes), or canvasbacks (A. valisneria). Stipkovits (1979) reported
pathogenicity of M. anatis to domestic ducklings and eggs, and neurological signs have
been recorded in ducks infected with M. anatis (Ivanics et al. 1988). Samuel et al. (1995)
infected game-farm mallard eggs with M. anatis and found reduced hatchling success,
hatchling size and growth rates. Hatchlings infected at 1 d of age did not have slower
growth rates. Goldberg et al. (1995) found esophagitis, tracheitis, and vaginitis in female
mallards from which M. anatis was isolated, and presented evidence for vertical
transmission ofM anatis in a wild gadwall (Anas strepera). Potentially, M anatis
infections could reduce recruitment in wild duck populations. Experimental infection of
ducklings withM gallisepticum resulted in suppressed growth rates (Stipkovits 1979).
Four other unidentified mycoplasmas were isolated from mallards, gadwalls, and black
ducks, but could not be associated definitively with disease (Goldberg et al. 1995).
Although M gallisepticum has been isolated from wild turkeys, most cases have
occurred in birds with close association to domestic poultry (Davidson et al. 1982, Jessup
et al. 1983, Luttrell et al. 1991, Fritz et al. 1992). Even though experimental infection of


ACKNOWLEDGMENTS
I thank Dr. Don Forrester for accepting me as a graduate student, and for his
support and guidance through the years. Drs. Kathy Ewel and Paul Gibbs assisted my
growth scientifically and professionally in the first half of my program. Dr. Ewels support
was instrumental in obtaining my fellowship, and Dr. Gibbs was responsible for a trip to
Australia. Drs. Mel Sunquist and Wiley Kitchens weathered the changes in my project
with grace and humor, and Dr. Kitchens was especially helpful in teaching me to argue my
positions and not back down when I knew I was right. It took several years for Dr. Mary
Brown to get me into her lab, and her support in presenting me to her colleagues is
appreciated. Dr. Elliott Jacobson has done his best to teach me clinical pathology and
histopathology and has been very supportive of my contributions to the overall project.
Dr. Paul Klein has given me some valuable insights into the critical thinking process.
Without my co-workers Drs. Dan Brown and Isabella Schumacher, Sylvia Tucker,
Barbara Crenshaw, and Cathie McKenna, and technicians Alyssa Whitemarsh, Michael
Lao, and Dave Bunger, this research would have been impossible. I benefited from Dans,
Isas and Barbs teaching abilities, and their willingness to discuss theory, practical
applications, philosophical underpinnings and differences of opinion. Mr. Clement
Lindsey and his staff cared for my research animals.
I thank Dr. Tim Gross for all his help with eggs and hatchlings, and John Wiebe
and Carla Weiser for their care. Drs. Dale Jackson and Michael Ewert provided advice.
Drs. Bruce Homer and Claus Buergelt have given me more strength and support than they
m


87
antibody levels, and it is unlikely that insufficient time had passed for the tortoises to
develop an immune response.
Environmental transmission of M gallisepticum is of considerable concern to the
poultry industry, necessitating disinfection of premises and equipment, and a 2-6 wk
fallow period before introducing new stock (Anonymous 1989, M. B. Brown pers.
comm.). The M. gallisepticum strain causing conjunctivitis in house finches also can
survive in the environment and cause infections in individuals later housed in the same
facility (Ley et al. 1996, Luttrell et al. 1996).
Environmental transmission of M agassizii in the wild may not be of great
concern. However, equipment used for capturing, handling, holding, and transporting
tortoises should be cleaned after each use by spraying or wiping with bleach, ethanol or
other disinfectant solution. Care should be taken to dispose properly of all gloves or
drapes that may have become contaminated. Clinically ill tortoises should not be housed
in direct contact with other animals, nor should indirect contact be allowed. Infected
tortoises should not be able to sneeze on other tortoises, nor should water or food dishes
be shared between pens without disinfection. Because the sample sizes at each time point
were very small, further research, with more rigorous attempts to isolate mycoplasmas and
quantify shedding, needs to be conducted. For the purposes of relocation, restocking, or
repatriation efforts, a short fallow period, perhaps 2 wk, is probably sufficient to prevent
environmental transmission of M. agassizii.


56
Therefore, the sentinel whose seroconversion was detected 8 wk after arrival probably
was exposed to M agassizii just before transport to UF.
Tortoises harboring M agassizii may not show clinical signs, may exhibit mild
signs, or may show signs only intermittently. Because tortoises not showing any ocular
signs or nasal discharge sometimes have positive culture and PCR results, the organism
may be transmissible from asymptomatic tortoises under appropriate conditions. Due to
unavoidable constraints on sampling live animals, I cannot exclude the possibility that
clinically healthy, seropositive, culture and PCR negative animals harbor the bacteria. As
shown by the two tortoises (the sentinel and the tortoise that was eliminated from the
study) that initially had negative culture and PCR results but developed disease without
being inoculated experimentally, some infections may go through an extended latent
period with low numbers of organisms in the nasal passages, or animals that appear to
have cleared the organism may, in fact, have not.
The clinical response of challenged animals was more rapid and severe than that of
naive animals, indicating that no protection was conferred by previous exposure to the
organism. This is consistent with some other mycoplasmal diseases in which the immune
response confers limited or no protection (Ellison et al. 1992), or contributes to
pathogenesis, such as arthritis in fowl caused by M. synoviae (Kume et al. 1977),
conjunctivitis in cattle caused by M. bovoculi (Rosenbusch 1987), and pneumonia in
humans caused by M. pneumoniae (Krause and Taylor-Robinson 1992). Tully et al.
(1995) have found that some mycoplasmal surface proteins share sequence and structural
homologies with vertebrate proteins, and suggest that these may play a role in eliciting
autoimmune responses. Repeated exposure to mycoplasmal proteins that resemble a


Table 5-1. Clinical signs, culture, PCR, and ELISA status of gopher tortoises included in the upper respiratory tract disease pairing
study at each sampling time. CS clinical signs (nasal or ocular discharge, palpebral edema, and/or conjunctivitis), Cl culture results.
P PCR results, E ELISA results.
Pen
ID
Aug. 1994
Oct. 1994
Mar. 1995
Aug. 1995
Feb.-Mar. 96
Jul.-Aug. 96
Necropsy
No.
Gender
CS Cl
p
E
CS
Cl
p
E
CS
Cl
P
E
CS
Cl
P
E
CS
Cl P E
CS P
E
Date
A1
226 F
Ra
201 M
-
R
A2
577 F
4-
sb
7/96
147 M
4-c
4-
4-
-
4-
- +
4-
+
7/96
B3
261 F
-
R
160 M
R
B4
129 F
-
IT1
211 M
R
B5
108 F
R
140 M
-
R
D1
135 F
4-
4-
-
s
4-
4- +
4-
4-
7/96
241 M
-
-
-
+
-
-
-
+
-
-
-
4-
+
4-
4-
4-
+
+ +
4-
4-
7/96
D2
275 F
+
-
-
+
-
-
-
+
-
-
-
4-
+
-
-
+
-
- 4-
-
+
7/96
144 M
disappeared
If
D3
151 F
4-
+
+
+
4-
4-
s
+
+
-
4-
+
-
-
S
4-
-
- -I-
S
+
7/96
126 M
-
-
-
-
4-
+
+
4-
4-
-
4-
4-
4-
4-
-
4-
+
- 4-
-
+
7/96
D4
186 F
S
-
+
4-
-
+
+
+ +
+
+
7/96
235 M
+
+
-
+
-
+
-
4-
+
-
-
4-
-
-
+
+
-
- 4-
s
4-
7/96
D5
213 F
-
-
-
-
+
-
-
4-
+
+
+
4-
4-
4-
4-
4-
euthanatised
6/95
185 M
+
-
4-
+
+
+
+
4-
+
4-
-
4-
+
4-
-
4-
euthanatised8
6/95
D6
150 F
-
-
-
-
+
+
+
S
4-
+
+
4-
4-
S
-
4-
4-
4- 4-
+
4-
7/96
183 M
+
4-
+
-
4-
+
-
+
+
+
+
+
4-
4-
-
4-
disappeared
Uh