The Etiology and pathogenesis of green turtle fibropapillomatosis


Material Information

The Etiology and pathogenesis of green turtle fibropapillomatosis
Physical Description:
xix, 284 leaves : ill. ; 29 cm.
Herbst, Lawrence Henry, 1957-
Publication Date:


Subjects / Keywords:
Turtles -- virology   ( mesh )
Neoplasms -- etiology   ( mesh )
Neoplasms -- veterinary   ( mesh )
Neoplasms, Experimental -- etiology   ( mesh )
Neoplasms, Experimental -- veterinary   ( mesh )
Herpesviridae -- pathogenicity   ( mesh )
Herpesviridae Infections -- etiology   ( mesh )
Herpesviridae Infections -- transmission   ( mesh )
Herpesviridae Infections -- veterinary   ( mesh )
Herpesviridae Infections -- physiopathology   ( mesh )
Trematoda -- pathogenicity   ( mesh )
Fibroblasts -- pathology   ( mesh )
bibliography   ( marcgt )
theses   ( marcgt )
non-fiction   ( marcgt )


Thesis (Ph. D.)--University of Florida, 1995.
Includes bibliographical references (leaves 258-283).
Statement of Responsibility:
by Lawrence Henry Herbst.
General Note:
General Note:

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Source Institution:
University of Florida
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All applicable rights reserved by the source institution and holding location.
Resource Identifier:
oclc - 49346555
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Full Text







Copyright 1995


Lawrence Henry Herbst

To my wife, Maria, and my two children, Thomas and Lisa,

who have supported me as I have pursued my dreams and who

have sacrificed the most to allow me that luxury. To my

parents who taught me the importance of family and the value

in doing what you love.


I want to express my deepest appreciation to Dr. Paul A.

Klein, chairman of my graduate committee, for the advice,

assistance, encouragement, support, patience, and friendship

that he has shown me throughout this project. I am also very

grateful to Dr. Elliott R. Jacobson, cochairman of my

graduate committee, for giving me the opportunity and

encouraging me to conduct this project. Special thanks also

go to Dr. Alan B. Bolten, Dr. Ammon B. Peck, Dr. Sheldon M.

Schuster, and Dr. John P. Sundberg, members of my graduate

committee, for their advice, assistance, and support.

A substantial part of this project would not have been

possible without the facilities and dedicated support

provided by Richie Moretti and Tina Brown of The Turtle

Hospital, Marathon, FL. Their sincere concern for the

environment and for endangered species has been amply

demonstrated by their commitment of time, labor, and money to

provide for the housing, husbandry, and rehabilitation of

marine turtles. I also thank veterinarians, Dr. Lisa Bramson

and Dr. Beatrice Lopez, for providing clinical care of

research turtles in my absence.

Many others have contributed in various ways to the

success of this research project. I especially thank George

Balazs (National Marine Fisheries Service, Southwest

Fisheries Laboratory, Honolulu, HI) for his continued

enthusiasm and interest in the fibropapilloma problem in

green turtles. Through his efforts, significant funding for

these studies were made available. I also thank Earl

Possardt, Ren Lohoefener, and Sandra McPherson of the U.S.

Fish & Wildlife Service for their support of this project.

The Florida Department of Environmental Protection (DEP)

provided permits to conduct these studies. In addition, the

cooperation and assistance of DEP personnel, especially

Barbara Schroeder, Allen Foley, Carrie Crady, and Blair

Witherington, in providing information about and access to

free-ranging or stranded turtles was extremely helpful.

Special thanks go to Dr. Llewellyn Ehrhart, University of

Central Florida, Orlando, FL, and his students and field

assistants, especially Dean Bagley and Steven Johnson, for

collecting blood, biopsy samples, and eggs. Eric Martin and

Barbara Schroeder also provided turtle eggs. Jim and Fern

Wood of Cayman Turtle Farm, Ltd, Grand Cayman, British West

Indies, made captive turtles available for antibody studies

and carried out immunizations. USAir Corporation transported

hatchlings from Gainesville to Marathon free of charge. Sea

World, Orlando, FL, provided access to turtles in their care

for blood sampling. Dr. Karen Bjorndal and Dr. Alan Bolten of

the Archie Carr Center for Sea Turtle Research provided

biological samples and allowed me to incubate eggs and house

hatchlings in their laboratory. Alan Bolten also provided

valuable assistance in negotiating the paperwork associated

with permits and work orders. I thank the personnel of the

Hybridoma Core Laboratory and Immunological Analysis Core,

Interdisciplinary Center for Biotechnology Research (ICBR),

especially Diane Duke, Linda Green, Cathy McKenna, and

Isabella Schumacher for their assistance. I also thank Dr.

Ratna Chakrabarty (Gene Expression Core, ICBR) for performing

the preliminary differential message display analysis. Dr.

Brian Gray (Cytogenetics Laboratory) karyotyped cell lines.

Dr. Ellis Greiner (Department of Pathobiology) sorted and

identified trematode parasites from green turtles. Dr. Bruce

Homer (Department of Pathobiology) allowed me to work at

times in his laboratory. Special thanks are due to Betty Hall

for teaching me immunohistochemistry techniques. Dr. Leonard

Shultz and Dr. John P. Sundberg (The Jackson Laboratory, Bar

Harbor, ME) provided immunodeficient mice for tumorigenicity


This project was supported by grants from SAVE-A-TURTLE,

Islamorada, FL, a joint contract from the U.S. Fish &

Wildlife Service, Department of the Interior and the National

Marine Fisheries Service, Southwest Fisheries Science Center,

NOAA, Department of Commerce (RWO No. 96), and a training

fellowship from the National Institutes of Health (National

Center for Research Resources RR07001).








. . . x ii

. . . x iv

. . . xv ii


Historical Perspective . .
Description of GTFP . .
Gross Pathology . .
Histopathology . .
Epizootiology of GTFP . .
Geographic Distribution . .
Prevalences . .
Seasonality . .
Demographic Patterns . .
Habitat Associations . .
Clinical Course, Morbidity, and Mortality
Possible Causes of GTFP . .
Environmental Factors . .
Ultraviolet light . .
Chemical contaminants .
Infectious Diseases . .

Viruses . .
Bacteria . .

Metazoan parasites . .
Genetic Factors .
Immune Dysfunction in GTFP Pathogenesis .
Conclusion . . .

TURTLE . . .


: : : : :


Introduction . .
Materials and Methods . .
Turtle Plasma Samples and Turtle
Immunizations . .
Preparation of Turtle Immunoglobulins
Hybridoma Production . .
Monoclonal Antibody Screening Protocols
Monoclonal Antibody Purification
and Biotinylation . .
Cross Species Reactivity of Monoclonal
Antibodies . .
Verification of Monoclonal Antibody and
Turtle Antibody Specificity .

Results . .
Immunoglobulin Purification .
Production and Characterization of
Monoclonal Antibodies .
Verification of Monoclonal Antibody
Specificity . .
Discussion . .


Introduction . .
Materials and Methods . .
Animal Housing and Maintenance .
First Transmission Study (1991) .
Materials for inoculation (1991)
Experimental turtles (1991)
Experimental treatments (1991)
Second Transmission Study (1992)
Material for inoculation (1992)
Experimental turtles and
treatments (1992) .
Third Transmission Study (1993) .
Material for inoculation (1993)
Experimental turtles (1993)
Experimental treatments (1993)
Histopathology . .
Transmission Electron Microscopy
Negative Staining Electron Microscopy
Results . .
First Transmission Study (1991) .
Second Transmission Study (1992)
Third Transmission Study (1993) .
Time to Tumor Development .
Histopathology of Experimentally
Induced Tumors . .
Transmission Electron Microscopy




. 87


S. 107

Negative Staining Electron Microscopy
Discussion . .
Experimental Evidence for an Infectious
Etiology . .
Role of Spirorchid Ova . .
The GTFP-Associated Herpesvirus .
Attempts to Identify Viruses in
Donor Material . .
Variation in Transmission Success .
Environmental Influences on Tumor

Development .
Conclusion . .

. 109
. 112

. 112
. 113
. 114

. 115
. 117

. 118
. 119


Introduction . .
Materials and Methods . .
Virus Isolation in Cell Culture .
Virus Extraction from Tumor Homogenates
Characterization Experiments .
Materials for inoculation .
Experimental turtles .
Transmission experiments with
tumor preparations .
Results . . .
Virus Culture . .
Virus Purification from Tumor
Homogenate . .
Characterization Experiments ..
Discussion . . .
Virus Culture . .
Virus Purification . .
Characterization Experiments .


Introduction . .
Materials and Methods . .
Histopathology . .
Immunohistochemistry for Detection of
Herpesvirus Antigens .
Immunofluorescence Staining for
Immunoglobulin Deposition .
Results . .
Cutaneous Fibropapillomas of Florida and
Hawaiian Turtles . .
Epidermal folding .
Pigmentation .
Pathologic changes in the dermis

. 120

. 120
. 121
. 121
. 124
. 124
. 127

. 128
. 129
. 129



. 143
. 144


. 148
. 148

S. 148
S. 149
. 152
S. 152

Pathologic changes in the epidermis 154
Inflammation . 163
Potential pathogens . .. 167
Experimentally Induced Cutaneous Tumors 173
Differences between spontaneous and
experimental tumors . 175
Early pathologic changes in
experimentally induced tumors 178
Herpesvirus Immunohistochemistry .. 180
Immunofluorescence for Antibody Deposition 181
Visceral Tumors . .. 181
Discussion . . 184
Florida Versus Hawaiian GTFP . 184
Spontaneous Versus Experimentally
Induced GTFP . 186
Pathogenesis of Tumor Progression .. 187
Pathogenesis of Dermal-Epidermal Clefts 190
Possible Routes of Dissemination .. 192
Can Herpesvirus Explain GTFP Pathology? 192
Can Papillomavirus Explain GTFP Pathology? .194


Introduction . . 196
Materials and Methods . .. 198
Plasma Samples . .... ..198
Detection of Antibodies to the
GFTP-Associated Herpesvirus .. 199
Detection of Antibody Reactivity
to Spirorchid Trematodes .. 201
Results . . 204
GTFP-Associated Herpesvirus .. 204
Spirorchidiasis . 207
Discussion . . 218
GTFP-Associated Herpesvirus .. 218
Spirorchidiasis . .. 220
Conclusion . . 225


Introduction . .. 226
Materials and Methods . .. 227
Cell Lines . . 227
In Vitro Assays .. . .. .. 229
Tumorigenicity Assay (In Vivo) 229
Detection of GTFP Cell-Associated Antigens 232

Detection of GTFP-Associated Changes in
Gene Expression . ... 233
Results . . 235
Cell Lines and In Vitro Characteristics 235
Tumorigenicity of GTFP-Derived Cell Lines 236
GTFP Cell-Associated Antigens .. 240
GTFP-Associated Changes in Gene Expression 244
Discussion . . 244


REFERENCES . . ... 258



Table 2-1. Prevalences of GTFP among different
populations of free-ranging green turtles 19

Table 3-1. List of monoclonal antibodies specific for
green turtle immunoglobulins . .. 72

Table 4-1. Free-ranging green turtles with cutaneous
fibropapillomatosis used as fibropapilloma donors 93

Table 4-2. Fibropapilloma development at inoculation
sites in green turtles treated with filtered cell-
free fibropapilloma extracts . .. 101

Table 4-3. Frequency of fibropapilloma development at
injection and scarification sites in recipient
green turtles . ... 104

Table 5-1. Free-ranging green turtles with cutaneous
fibropapillomatosis used as fibropapilloma donors 125

Table 5-2. Summary of characterization transmission
experiments . . ... 132

Table 5-3. The effect of chloroform treatment on the
infectivity of the GTFP agent ... .134

Table 5-4. Partitioning of GTFP agent infectivity by
ultracentrifugation . ... .135

Table 6-1. Comparison of the gross and histologic
features of fibropapilloma biopsies from free-
ranging Florida and Hawaiian green turtles .. 150

Table 6-2. Prevalence estimates of spirorchidiasis and
herpesvirus infection among green turtle population
samples . . 169

Table 6-3. Comparison of the gross and histologic
features of spontaneous and experimentally induced
fibropapillomas . ... 176


Table 7-1. Seroconversion to anti-herpesvirus
immunoreactivity among green turtles with
experimentally induced GTFP .

. 206

Table 7-2. Post-mortem diagnosis of trematode infections
in green turtles used as controls in ELISA
development . . 208

Table 8-1. Tumorigenicicty of fibroblast lines derived
from green turtles . .




Figure 2-1. Cutaneous fibropapillomatosis in the
green turtle, Chelonia mydas . .

Figure 2-2. Ocular fibropapillomatosis in green
turtles, Chelonia mda . .

Figure 2-3. Examples of visceral tumors found in
some green turtles with severe cutaneous
fibropapillomatosis . .

Figure 2-4. Variation in histologic appearance of
cutaneous fibropapillomatosis . .

. 7


Figure 2-5. Circumtropical distribution of green
turtle fibropapillomatosis (GTFP) .. 17

Figure 3-1. Fractionation of green turtle
immunoglobulins by gel filtration chromatography 67

Figure 3-2. Affinity purified turtle anti-DNP
antibody chains . . 69

Figure 3-3. Molecular masses of native 5.7S and
7S turtle immunoglobulins . ... 70

Figure 3-4. Sandwich ELISA demonstrating that putative
heavy chain specific Mabs bind plasma proteins with
immunoglobulin light chains . ... 74

Figure 3-5. Development of antibody responses to DNP
with time in 2 chronically immunized turtles

Figure 3-6. Inhibition of turtle anti-DNP antibody
activity by soluble antigen . .

Figure 4-1. Experimentally induced cutaneous
fibropapillomas in green turtles . .

Figure 4-2. Time course for experimental fibropapilloma
induction during the 1993 transmission study .






Figure 4-3. Experimentally induced fibropapilloma
showing characteristic benign epidermal hyperplasia
on broad fibrovascular stalks . .

Figure 4-4. Cytopathology in the epidermis of
experimentally induced green turtle
fibropapillomas . . .

Figure 4-5. Herpesvirus-like particles in experimentally
induced fibropapillomas . .

Figure 5-1. Subcellular particles identified in pooled
isopycnic gradient fractions prepared from
transmission-positive filtered tumor homogenate

Figure 6-1. Detection of herpesvirus antigens in green
turtle fibropapillomas by immunohistochemistry

Figure 6-2. Variation in the distribution of pigement

cells in cutaneous fibropapillomas

Figure 6-3. Typical cutaneous fibropapilloma in
the green turtle . .

Figure 6-4. Variation in the degree of epidermal
hyperplasia among cutaneous fibropapillomas

Figure 6-5. Degenerative changes observed in basal
epidermal cells in fibropapillomas .

Figure 6-6. Dermal-epidermal clefts in cutaneous
fibropapillomas . .

Figure 6-7. Degenerative changes observed in the
stratum spinosum of cutaneous fibropapillomas

Figure 6-8. Foreign body granulomas found in
fibropapillomas of free-ranging green turtles

Figure 6-9. Lymphocytic infiltrates observed in
green turtle fibropapillomas .

Figure 6-10. Herpesvirus-like intranuclear inclusions

Figure 6-11. Spirorchid trematode eggs found in
fibropapillomas of free-ranging green turtles

Figure 6-12. Bacteria, fungi, and algae found on
cutaneous fibropapillomas . .

Figure 6-13. Metazoan epibionts . .

. 155

. 156

. 158

. 159

S. 161

S. 164

. 166

S 168

. 170

. 172








. 153

Figure 6-14. Early pathologic changes in experimentally
induced fibropapillomas . .

Figure 6-15. Histologic features of visceral tumors
found in some green turtles with cutaneous GTFP

Figure 7-1. Spirorchid trematode ova recovered from
Florida green turtles . .

Figure 7-2. Plasma 7S IgY antibody responses of
controlturtles to crude adult spirorchid
antigen preparations . .

Figure 7-3. Relative frequencies of negative,
intermediate, and positive 7S IgY antibody
responses to Learedius learedii crude antigen
in turtle plasma samples from two sites .

Figure 7-4. Relative frequencies of GTFP negative
reef and lagoon green turtles having negative,
intermediate, and positive 7S IgY antibody
responses to Learedius learedii crude antigen
in plasma samples from GTFP-free turtles
from two sites . .

Figure 7-5. Relative frequencies of negative,
intermediate, and positive 7S IgY antibody
responses to Learedius learedii crude antigen in
plasma samples from GTFP-positive and GTFP-negative
Indian River lagoon turtles . .

Figure 8-1. Green turtle fibroblast cultures .

Figure 8-2. Tumorigenicity of GTFP-derived fibroblasts
in immunodeficient mice . .

Figure 8-3. Immunohistochemical detection of green
turtle fibroblasts in mouse ear fibromas .

Figure 8-4. Chromosomes of fibroblasts derived from
mouse ear fibromas . .

Figure 8-5. Differences in gene expression between
normal skin-derived and tumorigenic GTFP-derived
fibroblasts . . .






. 216

. 217







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



Lawrence Henry Herbst

December, 1995

Chairperson: Paul A. Klein
Major Department: Department of Pathobiology

Green turtle fibropapillomatosis (GTFP) is a threat to

populations of Chelonia mydas worldwide. This project

attempted to characterize the etiology and to describe the

pathogenesis of GTFP. Transmission studies showed that tumors

could be induced in recipient turtles by inoculation with

twice frozen and thawed cell-free homogenates prepared from

spontaneous tumors. Tumors were not induced by inoculation

with intact spirorchid ova nor were spirorchid ova found in

any experimentally induced tumors. Oncogenicity of tumor

homogenates passed through 0.45 pm but not 0.2 im filters,

and was destroyed by chloroform. Some spontaneous and

experimentally induced tumors had epidermal eosinophilic


intranuclear inclusions, which contained herpesvirus-like

particles. Attempts to culture this virus on 2 reptilian cell

lines were unsuccessful. Particles resembling herpesvirus

were found in pooled isopycnic gradient fractions of one

transmission-positive tumor preparation, but were not

tumorigenic. Green turtle antibody class-specific monoclonal

antibodies, developed for the detection of turtle antibody

responses to putative GTFP agents, were used with a proven

herpesvirus-specific turtle antiserum, to demonstrate

herpesvirus antigens in spontaneous and induced tumors.

Tissue sections containing herpesvirus were also used to

screen plasma samples for antibody reactivity to herpesvirus

antigens by immunohistochemistry. Antibody reactivities to

herpesvirus developed in all experimental transmission-

positive turtles, but not in controls or transmission-

negatives. A strong association between antibody reactivity

to herpesvirus and clinical GTFP was also found in free-

ranging turtles. In contrast, antibody reactivity to

spirorchid trematodes was not associated with clinical GTFP.

The transformed phenotype of GTFP-derived fibroblast cultures

was demonstrated using tumorigenicity assays and preliminary

studies showed differences in mRNA expression between matched

pairs of normal skin- and GTFP-derived cell lines. Although

the pathogenesis of GTFP can be explained by herpesvirus,

proof that herpesvirus causes GTFP will require reproduction

of the disease in turtles with purified virus, or


demonstration of herpesviral gene sequences among these

differentially expressed messages in GTFP cell lines and in

transmission positive tumor homogenates, that can transform

normal fibroblasts to the tumorigenic phenotype.



Reports of neoplasia in chelonians are relatively

uncommon (Billups & Harshbarger, 1976; Jacobson, 1980; 1981a;

Machotka, 1984). For example, 24 neoplastic conditions in 19

turtle species, mostly represented by single case reports,

were compiled by Machotka (1984). Cutaneous papillomas,

fibromas, and fibropapillomas in green turtles, Chelonia

mydas, however, were reported commonly (Machotka, 1984).

These three proliferative lesions in Chelonia mydas are the

hallmarks of Green Turtle Fibropapillomatosis (GTFP) and the

number of observed cases has continued to increase while only

2 new case reports of neoplasia in other turtle species have

subsequently been published (Frye et al., 1988; Machotka et

al., 1992). Recent documented increases in GTFP prevalence

and the spread of GTFP to locations where it had not been

observed previously make GTFP the most common neoplastic

disease of reptiles and a significant threat to endangered

green turtle populations. Consequently, research to determine

the cause of GTFP and find ways to reduce the impact of this

disease has been listed as a priority in recovery plans for

green turtles (Balazs et al., 1990; National Marine Fisheries

Service & U.S. Fish and Wildlife Service, 1991).

More recently, fibropapilloma-like lesions have been

reported in other marine turtle species, including loggerhead

turtles, Caretta caretta (Llewellyn Ehrhart, University of

Central Florida, Orlando, FL 32816, pers. comm.; Barbara

Schroeder, Florida Marine Research Institute, Tequesta, FL

33469, pers. comm.), olive ridley turtles, Leoidochelvs

olivacea (Any Chaves, Universidad de Costa Rica, San Jose,

Costa Rica, pers. comm.; Pamela Plotkin, Texas A&M

University, College Station, TX 77843, pers. comm.) and

flatback turtles, Natator depressus (Limpus & Miller, 1994),

raising concerns about the potential impact of these diseases

on all marine turtle populations.

The purpose of this research project has been to

identify the cause of GTFP, to lay the groundwork for

understanding the pathogenesis of this disease, and to begin

to develop practical diagnostic tests for use in management

applications and in ecological studies (epizootiology) of

GTFP. This research has addressed the important questions

that will lead to understanding of the impact of this disease

on worldwide green turtle populations.

The specific objectives were as follows:

(1) To review current knowledge about GTFP and evaluate

the various proposed hypotheses about its etiology (Chapter


(2) To begin to develop the immunological tools needed

to study the immune response to fibropapilloma cells or to

putative etiologic agents. The initial focus was to produce

monoclonal antibodies specific for the known classes of green

turtle immunoglobulins (Chapter 3).

(3) To determine whether GTFP can be transmitted

experimentally and is therefore caused by an infectious

(transmissible) agent (Chapter 4).

(4) To identify and characterize the etiologic agent

(Chapter 5).

(5) To rule out alternative infectious etiologies

(Chapters 4 & 5).

(6) To begin to describe the pathogenesis of GTFP based

on histopathology, experimental findings, and identified

serologic and epizootiologic associations (Chapters 6 & 7).

(7) To develop the techniques to distinguish cultured

GTFP-derived, i.e., transformed fibroblasts from normal

fibroblasts, as a basis for in vitro studies on the molecular

mechanisms of fibroblast proliferation, the most salient

feature of GTFP. In addition, to search for GTFP cell-

specific antigens or genes in these cell lines for use in

diagnostic test development (Chapter 8).


Historical Perspective

Cutaneous papillomas, fibromas, and fibropapillomas were

first described by Smith and Coates (1938) in a captive green

turtle, Chelonia mydas, at the New York Aquarium that had

been captured near Key West, Florida, two years previously.

Two other green turtles and 2 loggerheads that were housed

with this animal did not have lesions. Smith and Coates

(1938) also found fibropapillomas in 3 of 200 free-ranging

green turtles (27-91 kg) that were captured off of Key West.

That same year, Luck6 described similar tumors from a green

turtle caught off Cape Sable, Florida (Luck6, 1938). Masses

were located on the tail, flippers, axillae, neck, eyelids,

and corneas. Schlumberger and Luck6 (1948) subsequently

described fibropapillomas from 3 Florida green turtles and

found numerous fibrous masses within the lungs of one turtle.

In 1958, Hendrickson noted the occasional occurrence of

fibrous masses on nesting females in Sarawak and Malaya

(Hendrickson, 1958). The first confirmed case of GTFP in

Hawaii occurred in 1958 and was a juvenile green turtle

captured by local fisherman in Kaneohe Bay, Oahu (Balazs,

1991). A survey of local fishermen conducted by Balazs (1991)


suggests that GTFP was rare to nonexistent prior to this.

Since this first report, green turtles with fibropapillomas

have been reported with increasing frequency from Hawaii

(Balazs, 1991; George Balazs, National Marine Fisheries

Service, Southwest Fisheries Center, Honolulu, Hawaii 96822,

pers. comm.). In 1980 an outbreak of fibropapillomatosis

occurred in a breeding group of adult green turtles at Cayman

Turtle Farm, Ltd, Grand Cayman, British West Indies

(Jacobson, 1981b; Jacobson et al., 1989). The outbreak began

in wild caught adults but subsequently developed over several

years in farm raised turtles as well. Ehrhart (1991)

documented the first cases of GTFP in the Indian River

Lagoon, Florida, in 1982. Netting surveys within the northern

portion of the Indian River Lagoon system (Mosquito Lagoon)

had been conducted since 1977 without encountering any green

turtles with fibropapillomas. However, when the study area

was shifted to the central portion of the system (Indian

River) in 1982, affected turtles were encountered

immediately. A review of late 19th century accounts of the

Florida east coast green turtle fishery and of reports on

Indian River Lagoon turtles published between 1978 and 1983

failed to yield any record of GTFP prior to this (Ehrhart,

1991; Ehrhart et al., 1986). Continued monitoring at this

site since 1982 has revealed GTFP prevalences around 50%.

Description of GTFP
Gross Pathology

Green turtle fibropapillomatosis (GTFP) is characterized

by single to multiple raised cutaneous masses ranging from

0.1 cm to greater than 30 cm in diameter. Individual masses

may be either verrucous or smooth and either sessile or

pedunculated. Large masses are often ulcerated. Cutaneous

fibropapillomas are usually found on the soft skin but may be

found anywhere on the turtle's body, including carapace and

plastron. Common sites for GTFP are the flippers, neck, chin,

inguinal and axillary regions, and tail base (Figure 2-1).

Ocular GTFP is common, with masses arising from the bulbar

conjunctiva, limbus, cornea, or mucocutaneous junction of the

eyelids (Brooks et al., 1994; Jacobson et al., 1989; Luck6,

1938; Smith & Coates, 1938) (Figure 2-2). Tumor pigmentation

is usually related to the pigmentation of the skin at the

site of origin.

Visceral tumors (Figure 2-3) have been found at necropsy

in some green turtles with cutaneous fibropapillomatosis

(Jacobson et al., 1991; Norton et al., 1990; Schlumberger &

Luck4, 1948; Williams et al., 1994). Schlumberger and Luck6

(1948) discovered numerous spherical 3-5 cm masses in the

lungs of one green turtle. Norton et al. (1990) observed

multiple firm white nodules in both kidneys from a juvenile

green turtle with extensive cutaneous fibropapillomatosis

collected in the Florida Keys. Jacobson et al. (1991)

Figure 2-1. Cutaneous fibropapillomatosis in the green
turtle, Chelonia mvdas. This juvenile stranded in
December 1993 near Key West, Florida in severely
debilitated condition as evidenced by the sunken
plastron. Multiple tumors were found on the neck, front
and rear flippers, axillary and inguinal areas,
perineum, and covering both eyes.

Figure 2-2. Ocular fibropapillomatosis in green turtles,
Chelonia mvdas. (Top) Left eye with multiple
fibropapillomas originating from bulbar and palpebral
conjunctiva, limbus, and cornea. (Bottom) Right eye of a
second turtle with large fibromas arising from the
palpebral and bulbar conjunctiva.

Figure 2-3. Examples of visceral tumors found in some green
turtles with severe cutaneous fibropapillomatosis. (Top)
Kidneys with multiple, irregularly shaped, firm, white
nodules (arrows) ranging from 0.5 to 3 cm in diameter
bulging from the surface. (Bottom) Lungs with multiple
nodules ranging from 0.2 to 5 cm in diameter. Lung
nodules are well demarcated, smooth, and either firm and
white (fibromatous), or gelatinous and translucent
(myxomatous). Myxomatous nodules (arrow) appear to arise
from fibromatous nodules.

examined 2 turtles with GTFP and, in one animal, found

several discrete firm, white foci up to 1 mm diameter on the

surface of one kidney and multiple discrete 1-4 cm diameter

nodules in the other. They also found similar nodules 1-2 cm

diameter within both lungs. Williams et al. (1994) found lung

and kidney nodules in 41% (7 of 17) of the green turtles

examined from Puerto Rico. Approximately 17% (9 of 52) of the

green turtles with severe cutaneous fibropapillomatosis

presented for necropsy at a rehabilitation center have been

found to have similar nodules in the lungs, kidneys, and

other viscera (Herbst, pers. obs.; Richie Moretti & Tina

Brown, The Turtle Hospital, Marathon, FL 33050, pers. comm.).


Several histologic descriptions of cutaneous GTFP have

been published (Aguirre et al., 1994b; Brooks et al., 1994;

Harshbarger, 1991; Jacobson et al., 1989; Luck6, 1938; Norton

et al., 1990; Smith & Coates, 1938, 1939; Sclumberger &

Luck6, 1948; Williams et al., 1994). Cutaneous GTFP is

described as papillary epidermal hyperplasia supported on

broad fibrovascular stromal stalks (Figure 2-4). The ratio of

epidermal to dermal proliferation varies among lesions.

Masses in which both tissues are hyperplastic are termed

fibropapillomas (Figure 2-4A) while others, comprised of

proliferating dermal components with relatively normal

epidermis, are termed fibromas (Figure 2-4B). Several authors

have postulated that there is a developmental progression

from papilloma (early lesions) through fibropapilloma, to

fibroma (chronic lesions) (Harshbarger, 1991; Jacobson et

al., 1989; Luck6, 1938).

Varying degrees of orthokeratotic hyperkeratosis and

acanthosis were consistent features in all studies (Aguirre

et al., 1994b; Brooks et al., 1994; Harshbarger, 1991;

Jacobson et al., 1989; Luck4, 1938; Norton et al., 1990;

Smith & Coates, 1938, 1939; Schlumberger & Luck6, 1948). The

degree of epidermal hyperplasia in GTFP varied from mild to

moderate (7-15 cells thick) on skin tumors to extensive (up

to 30 cells thick) on some conjunctival and palpebral masses

(Brooks et al., 1994; Jacobson et al., 1989). Fibropapillomas

with extensive epithelial hyperplasia often exhibit

anastomosing rete ridges extending deep into the dermis.

Epithelial cells in hyperplastic areas tend to be

hypertrophied (Brooks et al., 1994; Jacobson et al., 1989).

The fibrovascular stroma contains numerous well-

differentiated fibroblasts arranged in a ground substance

containing compact bundles of collagen fibers. Fibroblasts

and collagen bundles tend to be haphazardly arranged, are

more numerous than in normal dermis, and are more dense near

the basement membrane (Brooks et al., 1994; Jacobson et al.,

1989). Various amounts of collagen and mucopolysaccharide


; :. .
S.- :..I ..
P' -' .

Figure 2-4. Variation in histologic appearance of cutaneous
fibropapillomatosis. (A) Fibropapilloma showing typical
arborizing pattern of papillary epidermal hyperplasia
supported by fibrovascular stroma.(B) Fibroma showing
extensive fibrovascular proliferation covered by
relatively normal epidermis. (H&E, scale bars = 250 pm).
relatively normal epidermis. (H&E, scale bars = 250 Gun).

ground substance have been demonstrated in cutaneous tumors

by trichrome and alcian blue staining (Norton et al., 1990).

Nerves and numerous small blood vessels are found within the

stroma. Fibropapillomas examined in several studies show no

malignant or anaplastic changes and few mitotic figures

(Smith & Coates, 1938; Williams et al., 1994). The benign

nature of GTFP has been confirmed by flow cytometry studies

(Papadi et al., 1995).

Some histologic features identified in cutaneous tumors

have not been reported consistently and may be incidental

findings. Trematode (Spirorchidae) eggs surrounded by

epithelioid macrophages and multinucleate giant cells were

found within the dermal capillaries of some fibropapillomas

(Aguirre et al., 1994b; Brooks et al., 1994; Harshbarger,

1991; Jacobson et al., 1989; Jacobson et al., 1991; Norton et

al., 1990; Smith & Coates, 1939; Williams et al., 1994). In

some lesions containing eggs, eosinophilic granulocyte

infiltrates were also observed (Smith & Coates, 1939).

Epithelial cells in the stratum spinosum and outer layers of

epidermis were hypertrophic and vacuolated in some GTFP

specimens. In these areas, amphophilic intranuclear

inclusions were sometimes observed (Aguirre et al., 1994b;

Jacobson et al., 1989). Jacobson et al. (1991) found

eosinophilic intranuclear inclusions containing herpesvirus-

like particles within some superficial epidermal cells

undergoing intracytoplasmic vacuolation and ballooning

degeneration. Lymphocytic perivascular infiltrates have been

described in several studies (Jacobson et al., 1989; Smith &

Coates, 1938). Cleft formation at the dermal-epidermal

junction has also been noted in fibropapillomas examined by

Jacobson et al. (1989).

Visceral tumors were composed of proliferating fibrous

tissue compatible with the dermal component of cutaneous GTFP

(Jacobson et al., 1991; Norton et al., 1990; Schlumberger &

Luck6, 1948). Lung nodules described by Schlumberger and

Luck6 (1948) were covered by ciliated columnar epithelium.

The renal nodules described by Norton et al. (1990) were

sharply demarcated from surrounding renal tissue and covered

by renal capsule at the surface. Normal renal tubules were

found scattered throughout the proliferating connective


Epizootiolorv of GTFP

Epizootiology is the study of the temporal and spatial

patterns of disease expression in animal populations and

includes efforts to identify etiology, describe incidence and

prevalence, morbidity and mortality, routes of natural

exposure or transmission, and the conditions that lead to

disease outbreaks epizooticss). This information is needed to

understand the full demographic impact of GTFP on wild turtle


Epizootiologic studies of GTFP have been hampered by

several factors. First, there are no diagnostic tests to

detect exposure and early (preclinical) disease because an

etiologic agent has not been identified. Thus, all prevalence

data are based on observation of gross cutaneous tumors

Second, because green turtles are migratory and long lived,

taking between 20 and 50 years to reach sexual maturity

(Balazs, 1982; Frazer & Ehrhart, 1985; National Marine

Fisheries & US Fish and Wildlife, 1991), it is difficult to

sample specific life history stages such as the post hatching

pelagic phase and impossible to conduct longitudinal studies

of cohorts. Third, attempts to correlate disease prevalence

with assorted biotic and abiotic factors are hampered by the

geographic scale over which field surveys need to be

conducted and by limited human and fiscal resources.

Consequently, surveillance for GTFP and monitoring of

potentially relevant biotic and abiotic factors has been


Even in well-monitored sites, sampling methods introduce

biases that affect prevalence estimates. Most field studies

are conducted on feeding grounds or nesting beaches, and it

is therefore not surprising that post pelagic juveniles and

adult females are over represented in the prevalence reports.

Nesting beach surveys underestimate the true prevalence of

GTFP in adult females because debilitated turtles are less

likely to nest and are therefore not sampled (Limpus &

Miller, 1994). Field surveys and fisheries that employ tangle

nets tend to selectively sample larger turtles because small

turtles are not caught in the mesh. Surveys based on stranded

sea turtles may overestimate the prevalence of severe

debilitating disease. Cold stunning events, and sampling

methods that use direct, in-water captures, provide the least

biased population samples. The reader is asked to keep these

caveats in mind when evaluating the information presented


Geographic Distribution

Presently, GTFP has a global circumtropical distribution

(Figure 2-5 and Table 2-1). GTFP has been reported from all

major oceans including the Atlantic (Florida, Bahamas,

Brazil), Caribbean (Cayman Islands, Puerto Rico, Virgin

Islands, Barbados, Venezuela, Colombia, Nicaragua, Costa

Rica, Panama, Belize), and Indo-Pacific (California, Hawaii,

Australia, Sri-Lanka, Seychelles, Sarawak, Malaya, Japan)

(Balazs, 1991; Balazs & Pooley, 1991; Gamache & Horrocks,

1991; Hendrickson, 1958; Jacobson, 1990; Jacobson et al.,

1989; MacDonald & Dutton, 1990; Limpus & Miller, 1994;

Williams et al., 1994; Karen Bjorndal & Alan Bolten,

University of Florida, Gainesville, FL 32611, pers. comm.;

Cynthia Lageaux, University of Florida, Gainesville, FL

32611, pers. comm.; Anne Meylan, Florida Marine Research

Institute, St. Petersburg, FL 33701, pers. comm.; Jean

Mortimer, University of Florida, Gainesville, FL 32611, pers.


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spread among regions. The early reports from Florida (Smith &

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the disease may have always had a worldwide, albeit sporadic



The prevalence of GTFP varies among locations and from

year to year. Table 2-1 summarizes the available prevalence

data from several field studies. The earliest published

prevalence estimate (1.5 %) was from a survey conducted in

1938 of turtles captured in the Key West, Florida fishery

(Smith & Coates, 1938). Most population surveys, however,

have been conducted since 1975. In these surveys, little or

no disease was found prior to 1982, but prevalences rose

rapidly in the 1980s and have remained elevated. Part of this

pattern may reflect an increased awareness of the disease,

but may also reflect a real increase in the prevalence and

severity of GTFP over time.

Prevalences in well monitored feeding ground sites range

from 0% in Inagua, Bahamas (Bjorndal & Bolten, pers. comm.),

Bermuda (Meylan, pers. comm.) and offshore reef sites in

Australia (Limpus & Miller, 1994), to 92% in Kaneohe Bay,

Hawaii (Balazs, 1991). Prevalences may vary greatly between

demographically matched populations over very short distances

(< 1 km), as seen when comparing the prevalence of GTFP in


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the Indian River (approximately 50%) with that from the

adjacent near-shore Sabellariid worm reef at Wabasso Beach

(0%) (Ehrhart, 1991; pers. comm.).


There is a seasonal pattern in the prevalence of GTFP

among stranded green turtles in Florida with more affected

turtles stranding in the winter months (Wendy Teas, Southeast

Fisheries Science Center, NMFS, Miami, FL, 33149, pers.

comm.). Anecdotal reports indicate that tumors grow rapidly

in summer and are quiescent in the winter in response to

water temperature (Moretti & Brown, pers. comm.). Thus,

tumors may grow rapidly in summer and may reach a size that

is debilitating by autumn. The onset of colder water

temperatures in winter may further stress GTFP affected

turtles sufficiently to cause the winter stranding peak.

Demographic Patterns

GTFP appears to effect certain age and size classes of

turtles more than others. GTFP is rare (0-12%) among nesting

adult females and lesions tend to be focal and mild (Balazs,

pers. comm.; Ehrhart, pers. comm.), although these data are

probably biased (see above). In Hawaiian feeding ground

sites, intermediate sized turtles, measuring 40-90 cm

straight carapace length (SCL), were more frequently and more

severely affected than other size classes (Balazs, 1991).

Ehrhart (1991) and Schroeder (pers. comm.) found similar

results in Indian River and Florida Bay, respectively.

Turtles weighing between 10 and 30 kg were more likely to

have GTFP and they were more severely affected than larger or

smaller size classes (Ehrhart, 1991). Similarly, among

stranded turtles in Florida, 93-98% are between 30 and 69.9

cm SCL (Teas, pers. comm.).

Juvenile green turtles enter near-shore feeding grounds

after 2-5 years of pelagic existence. Too few pelagic stage

juvenile green turtles have been examined to provide any

information about GTFP prevalence in this life history stage.

The consensus, however, is that juveniles develop GTFP after

they have migrated into near-shore waters. This hypothesis is

supported by the very low prevalence estimates for off-shore

sites (Table 2-1) and the fact that GTFP has never been

observed in any recent recruits from the pelagic environment

to near-shore habitats (Balazs, 1986; Ehrhart, 1991; Limpus &

Miller, 1994). Newly arrived juveniles are recognized by

their small size (25-30 cm SCL, weight < 5 kg) and the lack

of epibiota (algae, bryozoans, leeches, etc) on their

carapaces as seen in older resident turtles (Ehrhart, 1991;

Bolten & Bjorndal, 1992; Limpus & Miller, 1994). Although

these small turtles are under-represented in net surveys,

relatively unbiased samples of the near-shore juvenile turtle

population have been obtained during cold stunning events.

During one such event in 1985 in the Mosquito Lagoon,

Florida, 145 cold stunned turtles were collected

(Witherington & Ehrhart, 1985). Twenty-eight percent of these

turtles were recent recruits (< 5 kg) and none had GTFP even

though GTFP prevalence in the overall sample was 29%.

There are several explanations for the absence of

clinical GTFP among turtles that have recently immigrated to

near-shore feeding grounds from the pelagic environment. One

is that affected pelagic juveniles do not survive long enough

to migrate to feeding grounds. Survivorship of healthy

turtles through the 2-5 years of pelagic existence is already

so low that it is unlikely that diseased turtles would

survive. A second explanation is that the disease has a long

latent period, so that clinical disease develops only in

older juveniles after they have moved inshore, even though

exposure to the etiologic agents) may occur in the pelagic

zone. A third hypothesis is that the causative agents) or

environmental conditions appropriate for disease expression

are found only in some near-shore habitats, so that exposure

occurs after juveniles are recruited to these sites.

The wide variation in GTFP prevalence among size/age

matched populations of juvenile green turtles lends support

to the last hypothesis. If the cause of GTFP were encountered

in the pelagic zone and the pelagic juveniles assorted

randomly among near shore sites, then one would expect the

distribution of GTFP prevalences among these near-shore sites

to be more uniform than is observed. Monitoring studies in

the Indian River system provide the most convincing data

because GTFP prevalence varies from 0% to approximately 50%

over a very short distance (< 1 km) between two

demographically similar populations and there is documented

movement of turtles from the low prevalence site (ocean) into

the high prevalence site (lagoon) but not vice versa

(Ehrhart, 1991; pers. comm.).

Habitat Associations

The field survey data summarized in Table 2-1 indicate

that GTFP is more prevalent in shallow, near-shore ecosystems

(lagoons, bays) and possibly most prevalent in areas that are

impacted by human activities such as agricultural,

industrial, and urban development within the catchment basin.

The strongest association of GTFP prevalence appears to

be with habitat type (embayments). These marine environments

may provide favorable physical conditions for either

infectious or non-infectious disease agents. For example,

certain sediment types may accumulate chemical contaminants

and, combined with low flushing rates, could increase the

level of exposure to chemical carcinogens or immunotoxins.

These same sediment properties and hydrodynamic conditions

may also favor the accumulation and maintenance of high

concentrations of infectious agents. More variable water

temperatures in shallow embayments could affect the rate of

xenobiotic metabolism, tumor cell proliferation, immune

system function, and pathogen replication. For example,

thermal stress has been shown to exacerbate virus infection

in hatchling green turtles (Haines & Kleese, 1977; Kleese,

1984) and modulate virus expression and tumor growth in Luck6

renal adenocarcinoma (McKinnell, 1981, 1984; Zambernard &

Vatter, 1966). Variable salinity in near-shore habitats may

have similar stress effects.

Near-shore habitats may also provide an optimum biotic

environment for survival and transmission of an infectious

etiologic agent. Disease transmission could be enhanced by

high population densities of vectors or intermediate host

species. Feeding grounds may attract a high density of

susceptible turtles which would facilitate the transmission

of pathogens in a density dependent fashion, as has been

shown for horizontally transmitted damselfish

neurofibromatosis (Schmale, 1991) and the Luck6 virus

(McKinnell, 1981, 1984). Recruitment of susceptible turtles

from many different breeding stocks into common foraging

grounds may allow the exchange of many diseases, including

GTFP, from exposed to naive individuals. Habitat differences

in levels of other stressors such as concurrent infectious

disease (parasites), and disturbance by human activity

(fishing, boating, dredging) may render turtles more

susceptible to or less able to recover from GTFP. These

hypotheses provide the conceptual framework for future

epizootiologic studies.

Clinical Course. Morbidity, and Mortality

Accurate estimates of the number of turtles that become

clinically affected following exposure to the agents) that

cause GTFP are unavailable. The duration and course of

clinical GTFP are poorly understood, primarily because

individual turtles with fibropapillomas of known duration

have not been available for longitudinal studies. A few green

turtles have been held in captivity long enough to provide

some generalizations about clinical course of the disease.

Jacobson et al. (1989) held 6 immature turtles with multiple

cutaneous GTFP in captivity for several months. Some tumors

on some animals decreased in size while others increased in

some animals when examined 4 months after capture. Ehrhart

maintained 3 green turtles with GTFP in captivity for

approximately 3 months (Ehrhart, 1991; Ehrhart et al., 1986).

During that time one animal lost several tumors, a second

developed 8 new tumors, and the third exhibited no changes in

tumor burden. In these captive observation studies, the

length of time that animals had the disease prior to capture

was unknown.

Field mark and recapture studies also indicate a

variable clinical course. For example, of 56 green turtles

recaptured in the Indian River, 7% had tumors when first

marked but had none at recapture, 14% contracted tumors

between first capture and recapture, 38% had lesions both

times, while 41% were free of lesions both times (Ehrhart,

1991). These data, while limited in number, support the

conclusion that the clinical course is prolonged and that

some individuals may spontaneously recover from disease.

Recapture rates are generally low, however, and there is no

control over the time interval between capture and recapture.

Studies of the temporal patterns of progression and

regression of experimentally transmitted GTFP in captive

turtles are needed.

Accurate estimates of the proportion of clinically

affected turtles that die are unavailable. Cutaneous

fibropapillomas can become large enough to interfere with

locomotion and are easily entangled in discarded line. Ocular

fibropapillomas (Figure 2-2) may occlude vision and those

invading the cornea may cause secondary panophthalmitis with

destruction of the globe (Brooks et al., 1994; Herbst, pers.

obs.). Visceral fibromas (Figure 2-3) grow by expansion

within the stroma of the affected organ and eventually

disrupt normal organ functions. Cardiac dysfunction, buoyancy

problems and respiratory compromise, hydronephrosis,

gastrointestinal obstruction have all been observed or

suspected causes of death in affected turtles (Balazs, pers.

comm.; Herbst, pers. obs.; Moretti & Brown, pers. comm.).

Many green turtles with multiple cutaneous fibropapillomas

become severely debilitated (Figure 2-1). Blood chemistries

and blood cell counts of severely affected green turtles

confirm a general pattern of debilitation (Jacobson, 1987;

Norton et al., 1990). Abnormalities include non-regenerative

anemia, hypoproteinemia, electrolyte imbalances, uremia, and

elevations in liver enzymes (Norton et al., 1990). The

cachexia may be caused by any combination of the following:

inability to locate, ingest, or digest food, excessive energy

demands for growth by proliferating tumors, increased

energetic costs for locomotion, the physiological effects of

certain cytokines such as tumor necrosis factor, mediated by

the immune system, and/or concurrent disease such as

spirorchidiasis. Whatever the mechanismss, a number of

animals become sufficiently debilitated by GTFP to strand

(Balazs, 1991; Teas, 1991). In one rehabilitation center

about 50% of green turtles that were still alive at stranding

died despite extensive rehabilitation efforts (Moretti &

Brown, pers. comm.).

Possible Causes of GTFP

The etiology of GTFP is unknown and its identification

is one of the aims of this research project. Hypotheses about

etiology can be proposed based on comparisons with similar

tumors of known etiology from other species, the association

of potential pathogens with GTFP, and from epizootiologic

patterns. However, demonstration of causation requires

rigorous experimentation to fulfill Koch's postulates. This

is most easily accomplished with diseases caused by

infectious agents. However, infectious disease expression may

depend on a variety of host-related, pathogen-related, and

environmental factors (Hanson, 1988).

GTFP has histologic features in common with benign

cutaneous neoplasia, found in other vertebrates, such as

papillomas and fibromas (Pulley & Stannard, 1990), as well as

hyperplastic conditions such as keloidosis (Caro & Bronstein,

1985) and exuberant granulation tissue (Smith et al., 1972).

Thus, GTFP is consistent with either neoplasia or

hyperplasia. The classification of GTFP as a neoplastic

disease has been controversial (Harshbarger, 1984). In

general, neoplasia may result from any of a variety of

derangements at any point in the complex signalling and

control network of normal cellular proliferation and

differentiation (Bishop, 1991). The pathogenesis of neoplasia

may involve multiple cumulative steps (Hunter, 1991; Peraino

& Jones, 1989), with early steps causing unregulated

proliferation and later events leading to malignancy.

Similarly, hyperplasia can be caused by a variety of stimuli.

Thus, identifying a single etiology and fulfilling Koch's

postulates for GTFP may be difficult if not impossible.

Some potential causes of neoplastic and hyperplastic

proliferative lesions in other vertebrate species include

abiotic agents (ultraviolet light, chemical contaminants),

and infectious biological agents (viruses, bacteria, metazoan

parasites), with or without predisposing heritable genetic

conditions. The following sections review and discuss

evidence for or against the involvement of each of these

factors in the etiology and pathogenesis of GTFP.

Environmental Factors

Ultraviolet light

Smith and Coates (1938) were the first to suggest a role

for solar radiation in the pathogenesis of GTFP. Fifty years

later there is mounting concern that ozone depletion is

causing an increase in ultraviolet-B (290-320 nm) irradiation

(Kerr & McElroy, 1993) and that this may be having pervasive

effects in aquatic ecosystems (Hader, 1993) and on animal

health (Van der Leun & De Gruijl, 1993). UV-B produces direct

DNA damage by pyrimidine dimer formation (Anathaswamy &

Pierceall, 1990). This may lead to mutation in cellular

oncogenes and the development of neoplasia (Brash et al.,

1991). UV-B also causes immunosuppression in experimental

animals (Baadsgaard, 1991; Donawho & Kripke, 1991; Granstein,

1990; Noonan & DeFabo, 1992). The proposed mechanism involves

pyridine dimer formation (Applegate et al., 1989; Kripke et

al., 1992) and/or a trans to cis isomerization of urocanic

acid in the skin following UV-B absorption (DeFabo & Noonan,

1983; Noonan & DeFabo, 1992). For example, trout exposed to

levels of UV-B radiation within the ambient range recorded

for mid-latitudes developed skin damage and became

immunosuppressed, as evidenced by a high prevalence of fungal

skin infections (Fabacher et al., 1994).

Increased UV-B exposure could occur in the shallow

inshore waters where green turtles feed. However, GTFP

prevalence varies too greatly over very short distances (as

in Ehrhart's study area) for UV-B to be the cause of GTFP.

UV-B may be a cofactor in disease expression, however, and

the role of UV-B in modulating the immune system of turtles

deserves further investigation.

Chemical contaminants

A variety of chemical compounds have been shown to cause

benign fibroepithelial proliferation and to have mutagenic

and carcinogenic properties under experimental conditions

(Anderson & Reynolds, 1989; Weisburger, 1989). The list of

compounds is extensive but they seem to act by either of two

basic mechanisms: (1) direct nucleic acid damage leading to

genetic mutation initiatorss), and (2) cellular damage or

irritation leading to proliferation (promoters). As mentioned

earlier, chemical effects may be one of many mechanisms

involved in multistep carcinogenesis.

The involvement of chemical contaminants in naturally

occurring neoplastic disease of lower vertebrates has been

documented best in fish. The prevalence of liver pathology,

including liver neoplasia in brown bullheads, Ictalurus

nebulosus, was higher at contaminated sites than at

relatively clean sites in several North American lakes and

rivers (Baumann et al., 1987; Black, 1983; Bowser et al.,

1990a). Disease prevalence was correlated with contaminant

levels in fish in one study (Baumann et al., 1987) and with

sediment levels in another (Black, 1983). Neoplastic lesions

were induced experimentally by treating bullheads with

sediment extracts (Black, 1983). Similar associations between

hepatic neoplasia, polluted sites, and sediment contaminant

concentrations have been found for mummichogs, Fundulus

heteroclitus in Chesapeake Bay (Vogelbein et al., 1990) and

various bottom fish in Puget Sound (Malins et al., 1984). A

similar association has been found between contaminated

sites, in vitro mutagenesis of water and sediment extracts

from those sites, and the prevalence of pigment cell

neoplasia (Chromatophoromas) in croakers, Nibea mitsukurii

(Kimura et al., 1984; Kinae et al., 1990). In addition,

experimental application of chemical carcinogens reproduced

the tumors in these fish. In most of these studies,

polycyclic aromatic hydrocarbon (PAH) concentration was a

major factor in the association between disease prevalence

with contaminant levels. Similarly, PAHs have been implicated

in the pathogenesis of cutaneous neoplasia in tiger

salamanders, Ambvstoma tirrinum from a polluted pond (Rose,

1981; Rose & Harshbarger, 1977). Cutaneous papillomas have

been experimentally induced in lizards, Lacerta aailis with

dimethyl benzanthracene (Stolk, 1963).

Chemical contaminants may also play a role in the

pathogenesis of certain neoplastic diseases by disrupting

immune functions that would otherwise allow the host to

eliminate transformed cells. The effects of various

immunotoxins have been reviewed by Dean et al. (1990).

Chemical effects on immune function in fish has also been

reviewed (Anderson et al., 1984; Dunier, 1994; Zeeman &

Brindley, 1981). Associations between chemical contaminants

and immune dysfunction have been shown (Lahvis et al., 1995)

and experimentally demonstrated in some marine organisms

(Arkoosh et al., 1994; DeSwart et al., 1994). Contaminants

may also disrupt the immune system indirectly by disrupting

neuroendocrine functions (Colborn et al., 1993).

The role of chemical contaminants in green turtle

fibropapillomatosis is unknown. As described previously,

there is a possible association between high GTFP prevalence

and near-shore marine habitats that have been impacted by

human activity (Table 2-1). Although it is possible that

environmental degradation and contaminants play a role in

disease expression, more objective documentation of these

impacts in high and low GTFP prevalence sites are needed.

Problems arise in how to document the contaminant

exposure of marine turtles. Few data are available for

comparing contaminant residue levels in water, sediment, or

benthic organisms from high GTFP prevalence areas with those

from areas where GTFP is rare. Similarly, data on contaminant

levels in green turtle tissues are scant and difficult to

obtain because of the endangered status of this species

(Aguirre et al., 1994a; Clark & Krynitsky, 1980; Hall et al.,

1983; McKim & Johnson, 1983; Rybitski, 1993; Thompson et al.,

1974). The few studies that have been published are difficult

to interpret in the context of GTFP. For example, while one

study in 1983 found significant amounts of hydrocarbons in 2

green turtles that stranded after a major oil spill (Hall et

al., 1983), most surveys of organochlorine and

polychlorinated biphenyl residues in green turtle tissues

including egg (Aguirre et al., 1994a; Clark & Krynitsky,

1980; McKim & Johnson, 1983; Rybitski, 1993; Thompson et al.,

1974) have yielded relatively low levels, often below the

limits of detection of the methods.

Where data exist, there are problems with relating

contaminant levels to disease prevalence. First of all, the

biologic effect (toxicity) of any particular residue level in

green turtles is unknown. Second, surveys of residue levels

are usually limited to those chemicals that persist in the

environment or bioaccumulate, although important toxic

effects such as genetic damage (in a multistage

carcinogenesis model) can result from transient exposures to

compounds that do not bioaccumulate. In addition, exposure to

a potent chemical carcinogen may occur transiently in a

completely different habitat from that being monitored.

Third, toxic effects may not be direct as in some

experimental models, but may involve complex interactions

with other abiotic and biotic factors. Thus, fulfilling the

criteria for implicating chemical contaminants as the primary

cause of GTFP or as cofactor could be extremely difficult

(Foster et al., 1993; Hanson, 1988). Finally, the same

biological effects may be caused by any number of different

compounds acting through several different mechanisms.

Decisions about which contaminant residues to measure should

be made with specific a Driori mechanistic hypotheses in mind

or in light of documented history of exposure to specific


Nevertheless, there is a need to conduct further

toxicological studies. Specifically, there is a need to

collect data on the water, sediment, and turtle tissue levels

of several classes of chemical contaminants (including known

chemical carcinogens and immunotoxins) from several carefully

matched marine sites with different prevalences of disease.

In addition, controlled experiments involving exposure of

turtles to water or sediment extracts from high and low

prevalence areas will be necessary in order to clearly

demonstrate a contaminant effect in the etiology and

pathogenesis of this disease.

Infectious Diseases

The epizootiologic patterns observed among free-ranging

green turtle populations including the sudden appearance of

GTFP at new geographic sites, variation in prevalence over

relatively short distances, and temporal variation within a

locality are compatible with an infectious etiology. The

observation that some animals recover from GTFP is also

compatible with an infectious disease. In addition, an

infectious agent is the most plausible explanation for the

appearance and spread of GTFP among captive green turtles.

For example, the outbreak documented at Cayman Turtle Farm,

Grand Cayman, in 1980, began in wild caught adults and

subsequently developed in captive reared turtles over several

years. Once eliminated, GTFP has not recurred at Cayman

Turtle Farm despite little change in husbandry conditions

(Jacobson, 1981b; Jacobson et al., 1989). A similar outbreak

occurred in a head start facility in the Florida keys among 2

year old captive reared green turtles that had been held in a

pond where GTFP affected turtles were rehabilitated and

possibly had direct contact with affected turtles (Hoffman &

Wells, 1991).


A number of virus families (Papovaviridae,

Herpesviridae, Adenoviridae, Poxviridae, Retroviridae) are

known to induce proliferative and or neoplastic lesions.

Papillomaviruses (Papovaviridae) are the documented cause of

papillomas, fibromas, and fibropapillomas in many mammalian

and avian species (Sundberg, 1987) and are associated with

malignant neoplasia as well (Sundberg & O'Banion, 1989; Zur

Hausen, 1989). Among reptiles, a papillomavirus has been

described from hyperplastic skin lesions of 5 Bolivian side-

necked turtles, Platemvs Dlatvcephala (Jacobson et al.,

1982), and papovavirus-like particles have been observed in

papillomas of green lizards, Lacerta viridis (Cooper et al.,

1982; Raynaud & Adrian, 1976). A polyomavirus (Papovaviridae)

of hamsters produces benign cutaneous neoplasia in these

rodents (Graffi et al., 1968) but other polyomaviruses of

rodents and primates do not produce disease in their natural

hosts (Eckhart, 1990). Herpesviruses have been associated

with cutaneous papillomas and or fibromas in green lizards,

Lacerta viridis (Raynaud & Adrian, 1976), african elephants,

Loxodonta africana (Jacobson et al., 1986b), carp, CyDrinus

carpio (Hedrick et al 1990; Sano et al., 1985), and several

salmonids (Kimura et al., 1981a, 1981b, 1981c; Sano et al.,

1983; Yoshimizu et al., 1987). Poxviruses are responsible for

fibroepithelial proliferative lesions in squirrels (Hirth et

al., 1969; O'Connor et al., 1980), rabbits (Pulley & Shively,

1973; Shope, 1932), and primates (Behbehani et al., 1968).

Retroviruses have been associated with or proven to be the

cause of dermal sarcomas in walleyes, Stizostedion vitreum

(Martineau et al., 1991), lip fibromas in angelfish (Francis-

Floyd et al., 1993), neurofibromas in damselfish, Pomacentrus

partitus (Schmale & Hensley, 1988), myxofibromas and

rhabdomyosarcomas of snakes (Lunger et al., 1974; Zeigel &

Clark, 1969), sarcomas in poultry (Benjamin & Vogt, 1990),

and fibromas and sarcomas in a variety of mammalian species

including cats (Hardy, 1985; Moulton & Harvey, 1990) and

primates (Gardner & Marx, 1985; Tsai et al., 1990). The

molecular mechanisms of virus induced proliferation and

oncogenesis vary, but all in some way disrupt the cells'

signal transduction network (Benjamin & Vogt, 1990; Moran,

1993; Zur Hausen, 1991). Examples include, the E5 protein of

certain papillomaviruses that binds and activates the PDGF

receptor (Kulke & DiMaio, 1991; Petti & DiMaio, 1992; Petti

et al., 1991), the adenovirus ElB protein and papillomavirus

E6 protein that bind and inactivate cell cycle checkpoint

protein p53 (Benjamin & Vogt, 1990; Moran, 1993), and

adenovirus ElA, polyomavirus T antigen, and papillomavirus E7

proteins that target the cell cycle control protein pRB.

Certain viruses produce proteins that act like growth factors

or their receptors. For example, poxviruses may produce

epidermal growth factor (EGF)-like peptides (Brown et al.,

1985), simian sarcoma virus (a retrovirus) produces a PDGF-

like peptide (Benjamin & Vogt, 1990), and certain

herpesviruses express a protein with protein kinase activity

similar to receptor kinases (Smith et al., 1992).

Evidence for viruses. Certain histologic features of

GTFP, including perivascular lymphocytic infiltrates, vesicle

formation, and epithelial degeneration are consistent with,

although not specific for, virus infection, and have prompted

investigators to search for viruses. Smith and Coates (1938)

failed to find virus-like inclusions within the tumors that

they examined. Jacobson et al. (1989) examined tumors from

six turtles from Florida and one turtle from Hawaii by light

and electron microscopy. In some sections, cells in the

stratum spinosum and outer layers of the epidermis were

hypertrophic and vacuolated and amphophilic intranuclear

inclusion bodies suggestive of herpesvirus infection were

occasionally seen. Ultrastructural examination revealed mild

acanthosis (3-6 cells thick) and intracytoplasmic vacuoles

containing 150-170 nm diameter granules of varying electron

densities were described within the superficial epidermis but

not identified (Jacobson et al., 1989). Aguirre et al.

(1994b) described basophilic intranuclear inclusions in

several lesions that they suspected to be nucleoli, but also

considered compatible with viral inclusions. However, viral

particles were not found within these inclusions when

examined by EM. They also observed intracytoplasmic electron

dense granules approximately 150 nm in diameter that were

morphologically similar to viral particles. These, however,

were found in both normal and GTFP epithelium. These

intracytoplasmic granules are now generally accepted to be

mucin granules that are produced and secreted by normal

turtle keratinocytes as they differentiate (Aguirre et al.,

1994b; Jacobson, 1989; Matoltsy & Huszar, 1972).

Herpesvirus-like particles were demonstrated by electron

microscopy in some fibropapillomas taken from two juvenile

green turtles housed in the same rehabilitation facility in

the Florida Keys (Jacobson et al., 1991). In 3

fibropapillomas examined from one turtle and 1 of 14 tumors

examined from the second turtle, focal areas of ballooning

degeneration of superficial epithelium were found to contain

eosinophilic intranuclear inclusions. Electron microscopy

demonstrated the presence of particles within the nucleus

conforming in size and morphology (icosahedral 77-90 nm

diameter) with immature herpesvirus and intracytoplasmic

particles conforming with mature enveloped herpesvirus (110-

120 nm diameter). This agent was not successfully cultured so

experiments to fulfill Koch's postulates could not be


Immunologic and molecular methods have been used to

search specifically for papillomaviruses in GTFP biopsies

because of the similarities between turtle fibropapillomas

and those of other vertebrates known to be caused by this

type of virus (Sundberg, 1987). Jacobson et al. (1989) were

unable to demonstrate the presence of papillomavirus group-

specific structural antigens in paraffin embedded sections of

fibropapillomas from 1 Hawaiian and 6 Florida green turtles

using peroxidase-antiperoxidase immunohistochemistry. Total

DNA extracted from portions of these same tumors was probed

under low stringency conditions with bovine papillomavirus

type 2 virion DNA. Finally, a reverse Southern blot was

performed in which radiolabelled tumor DNA from two turtles

was allowed to hybridize with blots containing 25 different

cloned papillomavirus genomes (6 bovine, 7 human, and dog,

rabbit, coyote, mouse, rat, and parrot papillomaviruses).

These screenings for papillomavirus yielded negative results

(Jacobson et al., 1989). Similarly, Marc Van Ranst (Einstein

Medical College, Bronx, New York, pers. comm.) had negative

results when he performed low stringency southern blot

analysis on DNA extracts of 11 tumors collected from a single

immature green turtle from the Florida Keys. Probes included

full genomic DNA from human papillomaviruses HPV-1, HPV-2,

and HPV-5, bovine papillomavirus BPV-1, canine oral

papillomavirus, and pygmy chimpanzee papillomavirus PCPV-1.

Preliminary experiments have also been conducted using

the polymerase chain reaction and degenerate PCR primers for

conserved sequences in the El and L1 mammalian papilloma

virus genes. These primers failed to amplify any sequences in

11 GTFP biopsies from one green turtle (Van Ranst, pers.


These negative immunohistochemical and molecular data

are insufficient to rule out a papillomavirus as a potential

etiologic agent because papillomaviruses are extremely

diverse and it is not unlikely that a reptilian papilloma

virus would fail to react with mammalian and avian probes,

primers, and antisera (O'Banion et al., 1992).

The possible role of oncogenic retroviruses has never

been investigated. As with the papillomaviruses, retroviruses

may cause neoplasia without ever developing a patent life

cycle in the host (Benjamin & Vogt, 1990; Coffin, 1990).

However, in the absence of electron microscopic evidence for

virus production and shedding from the tumor it is difficult

to implicate a retrovirus as the cause. Detection of

integrated retroviral genomes (provirus) within the green

turtle genome will require specific molecular probes that

will become available only after the agent has been

identified and portions of its genome sequenced. Until then

it is unlikely that non-specific retroviral probes would

yield conclusive results given the ubiquity of endogenous

retroviral sequences in vertebrates (Coffin, 1990).


Chronic bacterial infections may induce proliferative

lesions in some tissues. For example, intracellular

Camovlobacter-like organisms are associated with

proliferative enteritis in ferrets, hamsters, and swine (Fox

& Lawson, 1988; Lawson et al., 1985). An invasive spirochaete

has been observed in papillomatous foot lesions in cattle but

experiments to fulfill Koch's postulates have not yet been

conducted (Read et al., 1992). Numerous species of bacteria

have been cultured from the surfaces of cutaneous green

turtle fibropapillomas (Aguirre et al., 1994b). However,

bacteria are not seen within intact GTFP lesions, and little

inflammation is observed within tumors unless the surface is

ulcerated, suggesting that these are all secondary

opportunistic pathogens.

Metazoan parasites

An association between parasites and neoplasia has been

made in several species. Dogs infected with the nematode,

Spirocerca lupi, which encysts in the esophagus, may develop

a fibrosarcoma at the site (Bailey, 1963). Rats with tapeworm

Cvsticercus (Taenia) cysts were reported to develop hepatic

carcinomas with high frequency (Dunning & Curtis, 1946).

Schistosoma mansoni infection with egg shedding in the

urinary tract has been associated with bladder cancer in

humans (Hashem et al., 1961).

Marine turtles are host to a variety of digenetic

trematode species (Lauckner, 1985). Benign papillomatous

lesions in the gallbladder of green turtles have been

associated with flukes and eggs of Rhvtidodoides similis

(family: Rhytidodidae) (Smith et al., 1941). At least 12

species of spirorchid trematodes have been described in the

green turtle (Lauckner, 1985). Their natural history is very

similar to Schistosoma in that adult trematodes inhabit the

vascular system and eggs must migrate through tissues to

reach an outlet to the environment. Schistosoma mansoni egg

antigens can elicit a fibrotic response in the host (Lammie

et al., 1986; Phillips & Lammie, 1986; Wyler, 1983) and it

has been suggested that spirorchid eggs may induce fibromas

by similar mechanisms (Harshbarger, 1984).

Evidence for a metazoan parasite etioloav. The

association of spirorchid egg deposition with

fibropapillomatosis was first noted by Smith and Coates

(1939), who found eggs of Hapalotrema constrictum in sections

of over half of 230 fibropapillomas that they examined from

Florida green turtles. Later, Jacobson et al. (1989) did not

find ova in any sections of 28 tumor biopsies collected from

6 Florida green turtles but eggs were present in tumor

sections from 1 Hawaiian turtle. Norton et al. (1990) and

Jacobson et al. (1991) found eggs in many sections of tumors

from 3 Florida turtles. Williams et al. (1994) found eggs in

fibropapillomas examined from 39 Caribbean green turtles, and

Aguirre et al. (1994b) found eggs in biopsy sections from 8

of 10 Hawaiian turtles affected with GTFP.

On the other hand, spirorchid ova have been found in 16

of 21 (76%) wild green turtles, 3 of 10 (33%) oceanarium-

reared green turtles, and 3 of 102 (2.9%) farmed green

turtles from Queensland, Australia where the prevalence of

fibropapillomatosis was 0% (Glazebrook & Campbell, 1990a,

1990b). In addition, ova and lesions associated with ova have

been found within otherwise normal tissues of GTFP affected

turtles (Aguirre et al., 1994b; Norton et al., 1990).

The significance of cardiovascular trematodes in the

etiology of GTFP remains unclear. On the one hand, some

authors have characterized cutaneous fibromas from green

turtles as a hyperplastic response to spirorchid eggs

(Harshbarger, 1984) although the occurrence of spirorchid ova

and ova induced lesions in otherwise normal tissues of

turtles with GTFP and in those that do not have GTFP argues

against a direct hyperplastic or tumorigenic effect. On the

other hand, Smith and Coates (1939) concluded that trematode

eggs were incidental, accumulating passively in the

microvasculature of tumors. Other authors have also tended to

discount the eggs as the cause of GTFP (Jacobson et al.,

1989; Lauckner, 1985).

Finally, an argument has been put forward that

ectoparasites may have some role in the pathogenesis of

fibropapillomatosis. Nigrelli (1942) and Nigrelli and Smith

(1943) found leeches, Ozobranchus branchiatus, infesting the

folds of papillomas and suggested that the leeches may act as

vectors of the causative agentss. Most authors agree,

however, that leeches do not cause tumors directly, although

they may severely debilitate their host (Schwartz, 1974). In

addition, hirudin secretion may cause some increased

vascularization at leech attachment sites (Lauckner, 1985;

Nigrelli & Smith, 1943). Aguirre et al. (1994b) reported

finding mites attached to the surface of Hawaiian

fibropapillomas but did not speculate on the significance of

this association.

Genetic Factors

Neoplastic transformation, by definition, involves

permanent, often multiple, changes in the cell's genotype

leading to relatively unregulated proliferation and

differentiation (Sirica, 1989). Familial patterns of

neoplastic disease arise from a heritable (germline) genetic

lesion that renders individuals more susceptible to disease

development following subsequent somatic cell genetic damage.

For example, a germ line loss of function mutation in a tumor

suppressor locus would predispose an individual to neoplasia

following any event that disables the remaining functional

allele (Haber & Housman, 1991; Knudson, 1986; Marshall,

1991). Familial patterns of neoplasia are well documented in

humans. Examples include Li-Fraumeni syndrome, Wilm's tumor,

retinoblastoma, and neurofibromatosis (Haber & Housman, 1991;

Marshall, 1991). Laboratory mice show extensive strain

variation in susceptibility to experimental tumorigenesis

(DiGiovanni, 1989). A well documented example of genetic

susceptibility to neoplasia among lower vertebrates is found

in certain platyfish, Xiphophorus maculatus/Xiphophorus

helleri hybrids, which have high rates of spontaneous and

ultraviolet light induced melanoma due to loss of a

functional tumor suppressor gene (Anders et al., 1984;

Friend, 1993; Sobel et al., 1975). Heritable defects in DNA

repair mechanisms could also render individuals more prone to

neoplasia as is the case in xeroderma pigmentosa (Dresler,

1989; Kraemer et al., 1984). The familial pattern of

epidermodysplasia verruciformis is believed to involve

heritable defects in cellular immune function and an

inability to eliminate papillomavirus infection (Orth, 1987;

Shah & Howley, 1990). In rabbits, certain major

histocompatibility loci have been associated with the

regression or progression to malignancy of Shope papilloma

induced tumors (Han et al., 1992). In addition, individuals

of some species have genetic predispositions to exuberant

hyperplastic responses to wounding, e.g. keloidosis in humans

(Caro & Bronstein, 1985) and "proud flesh" in horses (Smith

et al., 1972), that can resemble benign neoplasia.

The possibility that some green turtles have a genetic

predisposition to develop GTFP must be considered. However,

there is no evidence that this is the case because

genealogical studies in this species are impractical and

methods to distinguish susceptible from resistant individuals

are unavailable.

Immune Dysfunction in GTFP Pathoqenesis

The possibility that various biological and

environmental agents may be involved indirectly in GTFP by

causing immune system dysfunction has been alluded to in each

of the preceding sections. While immune suppression would not

be a necessary prerequisite for infection, if GTFP were

caused by a primary infectious agent, the disease would

probably be more persistent and severe (Chretien et al.,

1978; Duncan et al., 1975; McMichael 1967) and more likely to

progress to malignancy (Schneider et al., 1983) in those

individuals with compromised cellular immune function.

Implicating immune system dysfunction in the

pathogenesis of GTFP will be difficult because few turtle

specific reagents are available, immune function assays have

not been validated, and normal reference ranges have not been

established for green turtles. Immune system studies in sea

turtles are in their infancy. Collins (1983) provided initial

anatomic descriptions of green turtle lymphoid tissues. The

immunoglobulin classes of green turtles have been described

(Benedict & Pollard, 1972, 1977) and some preliminary

investigations of cellular immune functions have been

conducted (McKinney & Bentley, 1985). Matters are complicated

by the fact that the turtle immune system, as in other

poikilotherms, is influenced by both season and temperature

(Ambrosius, 1976; El Ridi et al., 1988; Muthukkaruppan et

al., 1982; Zapata et al., 1992). Eventually, systematic

surveys that compare immune function parameters among

apparently healthy turtles from populations with high and low

GTFP prevalences will be needed to test the hypothesis that

immune system dysfunction renders some green turtle

populations more susceptible to GTFP. If evidence for an

association between immune dysfunction and GTFP is found, it

will become important to identify those factors responsible

for immunomodulation in affected populations.


This chapter has brought together the available

published information about green turtle fibropapillomatosis

and provides the background for developing testable

hypotheses addressed in the following chapters of this

dissertation. It is clear from this review that the major

question to be addressed is, what is the etiology of GTFP?

Once the nature and identity of the etiologic agent are known

one can begin to develop strategies for monitoring and

preventing the spread of GTFP among turtle populations. With


techniques to monitor populations for exposure to the

causative agents) one can begin to model the long-term

demographic effects of this disease and initiate studies

designed to identify those factors that have allowed this

disease to become a worldwide epizootic.



There are 7 extant species of marine turtles, all of

which are threatened or endangered due to a variety of

factors such as over-harvesting, loss of nesting and feeding

habitats, marine pollution, and entanglement (National

Research Council, 1990). The impact of disease and the

relationship of disease susceptibility to environmental

factors are poorly understood, due in part to the lack of

diagnostic reagents with which to monitor the health status

of sea turtle populations. The importance of improved health

monitoring capabilities in wildlife conservation is becoming

increasingly recognized (Klein, 1993). The urgent need to

develop diagnostic tests for green turtles, Chelonia mydas,

stems in part from recent worldwide increases in the

prevalence and severity of green turtle fibropapillomatosis

and the need to better understand the epizootiology of this

disease (Chapter 2).

The development of standardized serodiagnostic tests for

green turtles would be facilitated by the availability of

monoclonal antibodies (Mabs) to specific turtle

immunoglobulin classes. Monoclonal antibodies are highly

specific and uniform reagents with reliable performance

characteristics that can be obtained in potentially unlimited

quantities. This paper describes the production and

validation of a battery of monoclonal antibodies specific for

each of the known immunoglobulin classes of the green turtle

(Benedict & Pollard, 1972).

Materials and Methods

Turtle Plasma Samples and Turtle Immunizations

Blood samples were collected into lithium heparin tubes

from the dorsal cervical sinus (Owens & Ruiz, 1980) of 4 green

turtles housed in a rehabilitation facility in Marathon,

Florida. The plasma obtained from these samples was used to

prepare immunoglobulins for use as antigen in hybridoma

production. Two juvenile captive-reared green turtles housed

at Cayman Turtle Farm, Grand Cayman, British West Indies were

immunized by biweekly subcutaneous inoculations with 250 tg

2,4-dinitrophenylated bovine serum albumin (DNP-BSA)

(Molecular Probes, Eugene, OR, USA) in Ribi's adjuvant (RIBI

ImmunoChem Research, Hamilton, MT, USA) for a total of 6

inoculations, followed by monthly inoculations of the same

DNP-BSA dose for another 8 months. A pre-immunization blood

sample was collected, followed by biweekly test bleedings

between each of the first 6 inoculations, and monthly

bleedings before each monthly booster inoculation. These

plasma samples were used to assess the ability of Mabs to

measure specific turtle anti-DNP responses and for affinity

purification of anti-DNP antibodies. In addition, pooled

plasma samples from loggerhead (Caretta caretta), olive

ridley (Lepidochelys olivacea), Kemp's ridley (LeDidochelvs

kempii), hawksbill (Eretmochelvs imbricata), and leatherback

(Dermochelvs coriacea) were obtained from various sources for

testing cross-species reactivity of the Mabs.

Preparation of Turtle Immunoalobulins

Several strategies were employed to isolate and purify

turtle immunoglobulins for use in mouse immunizations and

hybridoma screening. Initially, putative immunoglobulins were

identified and isolated according to their physico-chemical

properties. Later, additional approaches were taken, as

reagents and antigen specific antibodies from specifically

immunized turtles became available.

Globulins from a 50 ml sample of plasma from an

individual green turtle and from a 100 ml pooled sample from

3 green turtles from Marathon, Florida were precipitated with

saturated ammonium sulphate (SAS) (33% v/v ). The precipitate

was resuspended in PBS/az (0.01 M sodium phosphate buffer (pH

7.4) containing 0.15 M NaC1 and 0.02% NaN3) and the

precipitation repeated. The precipitate was dialyzed into

either PBS/az or 0.01 M Tris-HCl buffer (pH 8.0) and adjusted

to a final protein concentration of 2 mg/ml.

One portion (5 ml) of the globulin preparation (33% SAS

cut) in Tris buffer was applied to a diethylaminoethyl (DEAE)

anion exchange column and eluted in steps with 0.01 M Tris

buffer containing either 0.125 M NaC1, 0.25 M NaC1, 0.5 M

NaC1, or 1.0 M NaC1.

Another portion (18 ml) of the globulin preparation in

PBS/az was applied in 6 ml sample amounts to a 2.5 x 100 cm

Sephacryl S-300 column in order to separate proteins on the

basis of their size. Fractions were eluted with PBS/az at a

30 ml/hr flow rate and collected using a Gilson fraction

collector. Selected eluted protein fractions were reduced by

boiling for 5 minutes in Laemmli sample buffer (Laemmli,

1970) with 2-mercaptoethanol, and examined by SDS

polyacrilamide gel electrophoresis (SDS-PAGE) using a Phastgel

apparatus (Pharmacia LKB, Uppsala, Sweden). Fractions

containing similar protein composition were pooled and

concentrated in centrifuge filter concentrators (Amicon

CentriprepR-10, W.R. Grace & Co, Beverly, MA, USA). Selected

DEAE fractions were used to immunize mice and the gel

filtration fraction pools were used as antigen in preliminary

hybridoma screening protocols.

Immunoalobulin purification by anti-light chain affinity

column chromatographv. An affinity column was prepared using

2 mg monoclonal antibody HL673 which is specific for the

immunoglobulin light chain of the desert tortoise (Schumacher

et al., 1993). Mab HL673 which had been found to cross-react

strongly with putative light chain of the green turtle in

ELISA and Western Blots was covalently linked to a hydrazide

support gel (Affi-prepR Hz, Bio-Rad Laboratories, Richmond,

CA, USA) following manufacturers instructions. Briefly, 1 ml

of purified Mab HL673 (2 mg/ml) was oxidized with 20 pl of

sodium periodate stock solution (0.5 M NaI04) in oxidation

buffer (0.02 M sodium acetate, 0.15 M NaCl, pH 5.0) for 45

minutes at room temperature. The oxidation reaction was

stopped by addition of 5 p1 glycerol. The oxidized antibody

was dialyzed into coupling buffer (0.1 M sodium acetate, 1.0

M NaC1, pH 4.5) and incubated overnight with approximately 2

ml of settled hydrazide support beads. The antibody coupled

beads were then washed with 0.5 M NaC1, 0.01 M phosphate

buffer, pH 7.5 and stored at 40C. The column was prepared,

conditioned with elution buffer (0.1 M glycine, pH 2.7),

washed with PBS/az, and then 1 ml (2 mg) of green turtle

immunoglobulin rich preparation (33% SAS cut) was applied.

After washing the column the bound protein was eluted with

0.1 M glycine, pH 2.7. Fractions (1 ml) were collected and

neutralized with 45 p1 of 1.0 M Tris, pH 9.0. Eluted proteins

were concentrated and examined with SDS-PAGE. These proteins

were also used to immunize mice for hybridoma production.

Purification of anti-DNP antibodies by affinity column

chromatoaraphv. Turtle anti-DNP antibodies were purified

using affinity chromatography (Goetzl & Metzger, 1970; Wofsy

& Burr, 1969). N e-2,4-DNP-lysine (Sigma Chemical Co, St.

Louis, MO, USA) (2 mM in 0.1 M NaHCO3, pH 8.3) was coupled to

cyanogen bromide activated Sepharose 4B (Pharmacia LKB,

Uppsala, Sweden). The DNP-lysine coupled Sepharose was packed

into a column 2.5 x 13 cm (37 ml) and washed with 50 ml

methanol and then equilibrated in borate buffer (0.015 M

NaBO3, 0.15 M NaC1, pH 8.0). The column was further washed

with 1% bovine serum albumin (in PBS/az) and 25% acetic acid

followed by equilibration in high salt borate buffer (0.015 M

NaBO3, 0.5 M NaCi, pH 8.0) before use. Pooled plasma from the

two DNP-BSA immunized turtles was diluted 1:3 in high salt

borate buffer and applied to the column. The column was

washed until the optical density (OD28nm) returned to

baseline, and then any bound turtle anti-DNP antibodies were

eluted with 5 ml 0.1 M 2,4 DNP-glycine (pH 8.6). The eluted

fractions were pooled and concentrated to a 2 ml volume and

then extensively dialyzed against PBS/az. This solution,

containing highly purified turtle anti-DNP antibodies, as

judged by ELISA and SDS-PAGE, was used for final screening of

newly developed monoclonal antibodies. A small aliquot was

dialyzed against 50 mM Tris (pH 7.4) for mass


Mass spectrometrv. Affinity purified turtle anti-DNP

antibodies were submitted to the Protein Analysis Core,

Interdisciplinary Center for Biotechnology Research,

University of Florida for mass spectrometry (Vestec VT 2000,

Perseptive Biosystems/Vestec Mass Spectrometry Products,

Houston, TX, USA).

Hvbridoma Production

Mouse immunization protocols. One 6-8 week old female

BALB/c mouse was immunized subcutaneously with 6 Rg of HL673

affinity purified turtle immunoglobulin in Ribi's adjuvant.

Booster immunizations were repeated in two and four weeks.

The final booster was 17 ig of antigen intraperitoneally.

Fusion was performed 4 days after the last inoculation. Two

6-8 week old female BALB/c mice were immunized with a DEAE

fraction (50 gg total protein) containing both 5.7S and 7S

green turtle immunoglobulins (see results) in Ribi's adjuvant

at several subcutaneous sites. Booster immunizations were

performed at 2 weeks, 4 weeks (50 ig antigen per mouse).

Immunizations of both mice (100 gg antigen each) were

continued at 2 to 4 week intervals for a total of 7

immunizations using a combination of the 5.7S and 7S IgY rich

DEAE fraction (25-75 ig) and various IgM-rich preparations

derived from DEAE and Sephacryl S-300 chromatography runs

(45-100 gg). The two mice differed only in the last

immunization. One mouse was rested for about 4 weeks before

its final booster with both IgY and IgM whereas the second

mouse was rested for 10 weeks before its final booster with

turtle IgM alone. Serum anti-turtle titers were checked

periodically by ELISA.

Fusions. Monoclonal antibody production followed the

standard protocol of the Hybridoma Core Laboratory,

Interdisciplinary Center for Biotechnology Research,

University of Florida (Liddell & Cryer, 1991; Simrell &

Klein, 1979). Three independent fusions (one for each mouse)

were carried out. In general, four days following the final

booster immunization, mice were euthanized under

methoxyflurane anesthesia and their spleens removed.

Splenocytes were prepared by mechanical disaggregation,

washed, and fused with log phase SP2/0 mouse myeloma cells in

a 7:1 ratio using 50% polyethylene glycol 1500 media

(Boehringer Mannheim, Germany). After pelleting by

centrifugation at 400 x g for 8 minutes, cells were

resuspended in fusion media (D-MEM plus 1 x Antibiotic-

Antimycotic, 1 x HAT, 25% SP2/0 Conditioned Media, 20% Horse

Serum) (GIBCO, Grand Island, NY, USA) and seeded into 96 well

culture plates (Costar, Cambridge, MA, USA). Wells were

monitored microscopically for growth of hybridomas.

Screening was begun on growth positive wells 10-14 days

post fusion. The supernatants were removed and tested for

antibody reactivity against specific antigens (see below).

Hybridoma cultures of interest were transferred to 24 well

plates and expanded until they could be retested (about 7

days). Hybridoma cultures of interest were safeguarded by

cryopreservation in liquid nitrogen. Selected cultures were

isotyped using an isotyping kit (Amersham Mouse Monoclonal

Antibody Isotyping Kit, Code RPN.29, Amersham, UK) and cloned

by limiting dilution.

Monoclonal Antibody Screening Protocols

Hybridoma culture supernatants were screened against

each of the three turtle immunoglobulin rich pools derived

from the S-300 gel filtration column using enzyme linked

immunosorbent assays (ELISA). Secondary screening was done by

Western Blotting.

ELISA protocol. A standard ELISA protocol was used for

screening (Schumacher et al., 1993). Each well of a

microtiter plate (Maxisorp F96, NUNC, Kamstrup, Denmark) was

coated with 50 gl of antigen at a concentration of 10 gg/ml

in PBS/Az and incubated at 4C overnight. The wells were

washed four times with PBS/Az containing 0.05% Tween-20 (PBS-

Tween) by an automatic ELISA washer (EAW II, LT-Laboratories,

Salzburg, Austria) and then blocked with 300 il/well of 1%

BSA in PBS/Az at room temperature for 60 minutes or at 40C

overnight. After four more washes, 50 1l of hybridoma culture

supernatant was added to individual wells and incubated at

room temperature for 60 minutes. The wells were washed again

and 50 gl of a 1:1000 dilution of alkaline phosphatase

conjugated rabbit anti-mouse IgG whole molecule (Sigma

Chemical Co, St. Louis, MO, USA) was added to each well.

Following incubation at room temperature for 60 minutes, the

plates were washed 4 times with PBS-Tween and 100 gl of p-

nitrophenyl phosphate disodium (Sigma Chemical Co, St. Louis,

MO, USA) (1 mg/ml prepared in 0.01 M sodium bicarbonate

buffer, pH 9.6 containing 2 mM MgC12) was added to each well

and incubated in the dark at room temperature for 90 minutes.

The optical density of each well at a wavelength of 405 nm

was measured in an ELISA plate reader (EAR 400 AT, SLT-

Laboratories, Salzburg, Austria) at 30,60, and 90 minutes.

Positive and negative controls included on each assay plate

consisted of immune mouse serum and hybridoma cell culture

medium, respectively. Preliminary screens used selected gel

filtration fraction pools (either IgM-rich, 5.7S-rich, or 7S-

rich) as antigen (Figure 1). Later ELISA screens used

affinity purified turtle anti-DNP antibodies (2 gg/ml) as


Immunoblottina (Western blotting). Immunoblotting was

performed to help demonstrate the specificity of our

monoclonal antibodies for immunoglobulin chains. Immunoblots

were prepared following a published basic protocol

(Schumacher et al., 1993). Briefly, 100-150 gg of green

turtle globulins (33% SAS cut) were separated by SDS-PAGE

under reducing conditions, using a precast 10% Tris-glycine

gel (Novex, San Diego, CA, USA) as previously described

(Laemmli, 1970). The proteins were then electrophoretically

transferred from the gel to a nitrocellulose sheet (Schleicher

& Schuell, Keene, NH, USA) using a transfer apparatus (Novex,

San Diego, CA, USA). A Tris-glycine buffer (pH 8.3) in 20%

methanol was used as transfer buffer. Blotting time was 120

minutes at 30 volts. Once the transfer was complete, the

nitrocellulose was blocked immediately with 5% nonfat dry

milk in PBS/Az and incubated at room temperature on a rocker

overnight. The membrane was then washed three times (5

minutes per wash) with PBS-Tween and placed into a trough-

manifold (PR 150 Mini Deca Probe, Hoeffer Scientific

Instruments, San Francisco, CA, USA). Hybridoma culture

supernatants were loaded, 300 gl per channel, and incubated

on the nitrocellulose for 90 minutes at room temperature on a

rocker. The nitrocellulose membrane was washed 3 more times

and then incubated with 300 1l of a 1:1000 dilution of

alkaline phosphatase conjugated rabbit anti-mouse IgG whole

molecule for 90 minutes at room temperature. The membrane was

then removed from the manifold, washed 3 times and developed

with substrate buffer (0.1 M Tris-HCl, 1 mM MgC12, pH 8.8)

containing 44 p1 of nitroblue tetrazolium chloride (NBT) and

33 p1 of 5-bromo-4-chloro-3-indolylphosphate p-toluidine salt

(BCIP) per 10 ml of substrate buffer (Immunoselect, GIBCO

BRL, Gaithersburg, MD, USA). Immunoblots using biotinylated

Mabs followed the same basic procedure except that

biotinylated Mabs diluted in 1% BSA-PBS/az to 1 gg/ml

replaced hybridoma culture supernatants and strepavidin-

alkaline phosphatase (Zymed Laboratories, San Francisco, CA,

USA) replaced the alkaline phosphatase conjugated rabbit


Monoclonal Antibody Purification and Biotinvlation

Selected cloned hybridoma lines were injected i.p. into

Pristane-primed BALB/c mice and the resulting ascites fluid

containing the desired monoclonal antibodies was harvested.

Monoclonal antibodies were purified from ascites by passage

over a Protein G Sepaharose Fast Flow affinity column

(Pharmacia LKB, Uppsala, Sweden) and biotinylated.

Each purified Mab was dialyzed against 0.1 M NaHCO3, pH

8.0 and adjusted to a final concentration of 1.0 mg/ml

(Goding, 1986). Sulphosuccinimidyl-6-(biotinamido) hexanoate

(Immuno Pure NHS-LC Biotin, Pierce, Rockford, IL, USA)

dissolved in dimethyl sulphoxide at 1.0 mg/ml was added (120

gg biotin per mg of antibody) and the mixture was incubated

for 2 hours at room temperature. Following incubation, the

Mabs were dialyzed into PBS/az and stored at 40C.

Cross Species Reactivity of Monoclonal Antibodies

Supernatants from hybridomas producing Mabs specific for

green turtle immunoglobulins were screened by ELISA for

reactions with 33% SAS globulin preparations of 5 other sea

turtle species. The ELISA followed the general procedure but

used each 33% SAS cut at 5 pg/ml coating concentration as


Verification of Monoclonal Antibody and Turtle Antibody

The following experiments were conducted to prove

further that developed Mabs were specific for individual

turtle immunoglobulin classes and would react with turtle


Sandwich ELISA protocol. An antigen capture experiment

was designed to test whether the turtle plasma proteins bound

by each Mab possessed an immunoglobulin light chain. ELISA

plates were coated with 50 gl per well of selected purified

monoclonal antibody (5 gg/ml). Following incubation with

green turtle 33% SAS cut (2 gg/ml), the sandwich was

completed with 1 gg/ml biotinylated HL673 (anti-light chain)

and detected with strepavidin-alkaline phosphatase.

Detection of immune responses to DNP-BSA. Biotinylated

Mabs were used in an ELISA format to measure anti-DNP

antibody responses in turtles immunized with DNP-BSA. The

general ELISA protocol (described above) was used except that

Polysorp plates (Polysorp, NUNC, Kamstrup, Denmark) were

coated with 50 il per well DNP-BSA (1 gg/ml) and blocked with

2.5% casein (pH 7.0). Plasma samples from DNP-BSA immunized

turtles were diluted 1:50 in 1% BSA-PBS/az and serial two-

fold dilutions were tested. Plates were incubated with class

specific biotinylated monoclonal anti-turtle antibody (1

gg/ml in 1% BSA-PBS/az) followed by strepavidin-alkaline

phosphatase (Zymed Laboratories, San Fransisco, CA, USA).

Competitive inhibition assays. Competitive inhibition

ELISA's were used to verify that the turtle plasma proteins,

i.e. antibodies, detected by each Mab were DNP-specific.

First, plasma samples with peak anti-DNP responses from the 2

immunized turtles were serially diluted and incubated at 4C

overnight with increasing concentrations of soluble hapten

(2,4 DNP-glycine pH 7.4 in PBS/az final concentration range:

0-2 mM). These "inhibited" plasma samples were then assayed

for residual antibody activity by ELISA as described above.

Second, plasma samples with peak anti-DNP responses were

serially diluted and mixed with serial twofold dilutions of

rabbit anti-DNP antiserum (Sigma Chemical Co., St. Louis, MO)

with specific anti-DNP antibody concentrations ranging from 0

to 7.5 gg/ml. The residual turtle DNP-specific antibody

activity was then assayed by ELISA.


Immunoclobulin Purification

Turtle globulins eluted from the DEAE column in two

peaks corresponding to 0.125 M NaC1 and 0.25 M NaC1. Analysis

by reducing SDS-PAGE revealed that the 0.125 M NaCl peak

contained three major protein components with approximate

molecular weights of 23 kD, 38 kD, and 65 kD, consistent with

immunoglobulin light chain, 5.7S heavy, and 7S heavy chains

respectively (see Ambrosius, 1976). The 0.25 M peak contained

a mixture of proteins of various sizes, including a 70 kD

band consistent with IgM heavy chain (data not shown).

Turtle globulins separated on the Sephacryl S-300 column

also eluted in two major peaks: an early peak containing

putative IgM and a large late peak containing a mixture of

5.7S and 7S IgY (Figure 3-1A). Fractions were analyzed by

reducing SDS-PAGE and those with similar protein composition

were pooled and the resulting pools were designated as either

IgM-rich, 5.7S-rich, or 7S-rich (Figure 3-1B). These were

used for initial ELISA screening of hybridoma supernatants.

A very small amount of turtle immunoglobulin (< 100 pg)

was purified from the globulin preparation with an anti-light

chain immunoaffinity column. This material contained three

major components of approximately 65 kD, 38 kD, and 23 kD,

consistent in size with turtle 7S and 5.7S heavy chains and

light chain (data not shown).

Turtle anti-DNP antibodies eluted from the DNP-Sepharose

column showed 4 predominant bands on reducing SDS-PAGE. These

bands had molecular weights of approximately 23, 38, 65, and

70 kD as expected of the light chain, 5.7S, 7S, and IgM heavy

chains respectively (Figure 3-2A). This protein was used in

ELISA and western blotting to screen the monoclonal antibody

supernatants and purified biotinylated Mabs. Figure 3-2B

shows representative western blot results for these Mabs and

HL673 (anti-tortoise light chain) and demonstrates the

specificity of each Mab for its target immunoglobulin chain.

The control Mab, HL860 (anti-turtle non-immunoglobulin), did

not react with the affinity purified turtle anti-DNP antibody


An aliquot of affinity purified turtle anti-DNP

antibodies was examined by mass spectrometry. The mass

spectrometer detected two proteins with molecular weights of

120 and 175 kD corresponding to the expected molecular

weights of intact 5.7S and 7S Ig respectively (Figure 3-3).

Figure 3-1. Fractionation of green turtle
immunoglobulins by gel filtration chromatography.
Turtle globulins (33% SAS cut) were applied to a
Sephacryl S-300 column and eluted with PBS/az.
Fractions with similar protein composition by SDS-
PAGE analysis were pooled. Three fraction pools
were produced: IgM rich (fractions 8-18), 7S rich
(fractions 34-40), and 5.7S rich (fractions 50-62).
(A) Elution profile of turtle globulins
fractionated on S-300 column. Protein content of
each fraction was estimated by spectrometry
(OD28onm). (B) Reducing SDS-PAGE (10% Tris-glycine)
of green turtle immunoglobulin rich fraction pools,
stained with Coomassie Blue. Lane 1--molecular
weight markers (kD); lane 2--33% SAS cut; lane 3--
IgM rich pool; lane 4--7S IgY rich pool; lane 5--
5.7S IgY rich pool.

0 20 40 s0


1 2 3 4 5

97 p -
69 > <
46 -w

U -

Amm --


1 2 1 2 3 4 5 6 7
< 200

S 974 4 974
S69 4 69
OW 46
< 46

N 4 30 b

l4 21.5 I4 30

4 21.5

Figure 3-2. Affinity purified turtle anti-DNP antibody
chains. (A) Coomassie Blue stained 12% Tris-glycine
reducing gel showing turtle anti-DNP antibodies eluted
from a DNP-Sepharose affinity column with 0.1 M DNP-
glycine. Lane 1--turtle anti-DNP antibodies; lane 2--
molecular weight markers. (B) Immunoblot of selected
monoclonal antibodies (Mabs) on affinity purified turtle
anti-DNP antibodies. Each lane was incubated with a
different biotinylated Mab: either HL860 anti-turtle
non-immunoglobulin plasma protein (lane 1), HL846 anti
IgM (lane 2), HL857 anti-7S IgY heavy chain (lane 3),
HL814 anti-5.7S IgY heavy chain (lane 4), or HL673 anti-
tortoise IgY light chain (lane 5). Control lane
contained 1% BSA (lane 6). Lane 7 contained molecular
weight markers.

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Native IgM could not be detected by mass spectrophotometry

because the size limit for the method is 200 kD.

Production and Characterization of Monoclonal Antibodies

Hybridoma screening by ELISA against various

immunoglobulin rich fraction pools and Western blotting

against turtle globulins (33% SAS cut) yielded 20 hybridomas

of interest which were retained for further study. The

initial selection of these hybridomas was based on the

specificity of their Mabs for turtle proteins of the

appropriate physical and chemical properties. However these

results were not sufficient proof that these Mabs were

specific for turtle antibodies. Further screening by ELISA

against affinity purified turtle anti-DNP antibodies showed

that only 15 were specific for turtle immunoglobulin classes.

Table 3-1 gives the specificities and isotypes of these 15

Mabs. Ten of these Mabs were specific for 7S IgY heavy chain,

whereas 2 Mabs each were specific for the immunoglobulin

light chain and IgM heavy chain, and only 1 Mab was specific

for the 5.7S IgY heavy chain.

Mabs from the 15 hybridomas that reacted positively with

affinity purified turtle anti-DNP antibodies were tested by

ELISA against serum globulin fractions (33% SAS cuts) from 5

other sea turtle species. Table 3-1 shows that several

monoclonal antibodies reacted with epitopes that are shared

broadly among sea turtle species. Nine of the 7s IgY heavy

chain specific Mabs cross-reacted with all sea turtle


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species. Both light chain specific Mabs cross-reacted with

all species except the leatherback turtle. The IgM and 5.7S

Mabs on the other hand seemed to be specific for green turtle


Verification of Monoclonal Antibody Specificitv

Three hybridomas were cloned and their monoclonal

antibodies purified. These were designated HL814 (anti-5.7S

IgY heavy chain), HL846 (anti-IgM heavy chain), and HL857

(anti-7S IgY heavy chain) respectively. These Mabs were used

for further validation experiments. Purified Mabs against

turtle immunoglobulin light chain were not prepared because

of the availability of HL673 (anti-desert tortoise light


Sandwich ELISA. An experiment was designed to test

whether the turtle plasma proteins being bound by each of

these Mabs could be identified as immunoglobulin by the

criterion of having an immunoglobulin light chain. Figure 3-4

shows that proteins, selectively captured from turtle plasma

SAS cut by Mabs HL814, HL846, or HL857, in turn bound

labelled light chain specific Mab HL673, whereas antigen

captured by HL860 (Mab specific for an unidentified 33 kD

turtle protein present in SAS cut) failed to bind Mab HL673.

Antibody Responses to Immunization. Figure 3-5A-D shows

that both HL814 and HL857 could detect rising anti-DNP

antibody responses in both experimentally immunized turtles.

A rise in 7S IgY anti-DNP activity was detected in both





0 0 -

HL814 HL857 HL846 HL860 PBS


Figure 3-4. Sandwich ELISA demonstrating that putative heavy
chain specific Mabs bind plasma proteins with
immunoglobulin light chains. ELISA plates were coated
with 5 pg/ml of each of the following purified Mabs:
HL814 (anti-5.7S heavy chain), HL857 (anti-7S heavy
chain), HL846 (anti-IgM heavy chain), and HL860 (anti-
non-immunoglobulin plasma protein). Following incubation
with 2 gg/ml turtle globulins (33% SAS cut), plates were
washed and incubated with 1 gg/ml biotinylated HL673
(anti-tortoise light chain) to complete the sandwich.
Binding of HL673 was detected with strepavidin alkaline
phosphatase. Data presented are ELISA reactivities
(OD405nm) after 90 minutes incubation with substrate.

turtles within 5 weeks of beginning immunizations and

remained high for the remainder of the experiment. The rise

in 5.7S IgY anti-DNP activity took up to 9 months to reach a

maximum in both turtles. Results for IgM (HL846) were less

clear. One turtle (#4624) showed a weak IgM peak at about 13

weeks (Figure 3-5F), whereas the other turtle (#3150)

appeared to have a high IgM anti-DNP response in the pre-

inoculation sample as well as subsequent samples (Figure 3-

5E). Various modifications of the ELISA protocol, such as

using high salt (0.5 M NaCl) buffer, failed to reduce the

pre-inoculation putative IgM anti-DNP signal. Neither turtle

developed detectable antibody titers to BSA after 10 months

of immunization with DNP-BSA.

Inhibition by soluble hapten. Figure 3-6 (A-C) shows

that the ELISA reactions of immune plasma having peak anti-

DNP antibody titers can be inhibited with increasing

concentrations of soluble hapten (DNP-glycine). Both the

degree of inhibition attained and the shape of the inhibition

curves varied between turtles and among antibody classes

assayed. Inhibition ranged from 72 to 97% with 1 pM DNP-

glycine. Because no anti-BSA antibody responses could be

detected in either turtle, it was impossible to test whether

peak anti-BSA responses would be refractory to inhibition by

soluble DNP-glycine. Nevertheless, results of this experiment

support the conclusion that these Mabs (HL814, HL846, and

HL857) recognize DNP-specific antibodies. The ELISA reactions

Figure 3-5. Development of antibody responses to DNP
with time in 2 chronically immunized turtles.
Serial two-fold dilutions of plasma samples,
collected periodically throughout a prolonged
immunization schedule (10 months), were tested by
ELISA for anti-DNP activity using various
biotinylated Mabs. The rise in OD405nm with time is
indicative of a rising DNP-specific antibody titer.
Turtle 7S IgY (A & B), 5.7S IgY (C & D), and IgM (E
& F) responses were detected by biotinylated HL857,
HL814, and HL846 respectively. Panels A, C, and E
show the responses of turtle #3150 and B, D, and F
show those of turtle #4624.



Dilution 1:Po


1 :2
Dilution 1:3m0




1:50 2
1:200 :
Dilution 1:3

1:Di 0o
Dilution 1:SlO



Dilution 0
Dilution 1:32oo

r 33
I-- =

Time (weeks)

Figure 3-6. Inhibition of turtle anti-DNP antibody
activity by soluble antigen. Serially diluted
plasma samples with peak 5.7S, 7S, or IgM anti-DNP
antibody activity were incubated overnight with
final concentrations of 2,4 DNP-glycine ranging
from 0 to 1 pM. Samples were then tested by ELISA
for binding to DNP-BSA coated plates. Readings
(OD405n) taken at 60 minutes were plotted against
DNP-glycine concentration. The plasma samples used
were: week 41 for 5.7S and 7S (both turtles), week
1 for #3150 IgM, and week 13 for #4624 IgM (see
Figure 3-5). Data presented are for plasma diluted
1:400 for 5.7S IgY (HL814) and 7S IgY (HL857) and
diluted 1:50 for IgM (HL846). Solid bars (turtle
#3150); crosshatched bars (turtle #4624).



? 2



0 10 40 100 200 S00 1000

o B
S 2



0 10 40 100 200 500 1000







0 10 40 100 200 500 1000



of immune plasma with peak anti-DNP antibody titers were also

strongly inhibited by increasing concentrations of rabbit

anti-DNP specific antibodies (data not shown).


Sea turtles have 3 major classes of immunoglobulins: a

17 S IgM, a 7S IgY, and a 5.7S IgY (Benedict & Pollard,

1972). IgM is believed to be produced transiently early in an

immune response, as in mammals (Benedict & Pollard, 1977;

Chartrand et al., 1971)- In reptiles, IgM may be the primary

immunoglobulin that is secreted onto mucosal surfaces (Portis

& Coe, 1975). The 7S IgY is believed to function as a serum

antibody like mammalian IgG. The role of 5.7S IgY is unclear,

but evidence suggests that it is a chronic immune response

globulin and that it is maternally transferred to egg yolk

(Benedict & Pollard, 1972, 1977; Chartrand et al., 1971).

The results presented here show that Mabs with

specificity for the light chain and each of the three heavy

chain classes of the green turtle have been produced. The

purification and screening strategies used were dictated in

part by the limited availability of turtles and antigen

specific plasma. Initially, plasma from specifically

immunized green turtles was unavailable, so the preliminary

immunoglobulin purification and hybridoma screening relied on

identification of plasma proteins with the physico-chemical

properties (solubility, size, and charge) consistent with

earlier reports on turtle immunoglobulins (Benedict &

Pollard, 1972, 1977; Leslie & Clem, 1972). Previous

structural studies of turtle antibodies (several species)

indicated that turtle light chains are approximately 22.5 kD

and turtle 5.7S IgY, 7S IgY, and IgM heavy chains are 35-38,

63-68 kD, and 70 kD respectively (Ambrosius, 1976; Benedict &

Pollard, 1977; Chartrand et al., 1971; Leslie & Clem, 1972).

Preliminary screening of fusions yielded a collection of 20

hybridomas that bound to turtle plasma proteins with the

appropriate physico-chemical properties. However, additional

screenings against antigen-specific turtle antibodies

(affinity purified turtle anti-DNP antibodies) revealed that

only 15 of these Mabs could be classified as immunoglobulin

specific. The turtle proteins recognized by the other 5 Mabs

had similar physico-chemical properties but could not be

shown to bind antigen.

Additional experiments were conducted to prove further

that the selected cloned hybridomas produced Mabs that were

specific for turtle antibodies. A sandwich ELISA, using an

anti-light chain Mab demonstrated that the turtle plasma

proteins recognized by each of the heavy chain specific Mabs

possessed a light chain, thereby confirming the specificity

of these Mabs for turtle immunoglobulins. Mabs specific for

turtle immunoglobulin classes should be able to detect an

increasing antibody titer in response to immunization with

specific antigen. Mabs HL814 and HL857 (5.7S IgY heavy and 7S

IgY heavy chain specific, respectively) were able to measure