PIROPLASMS IN FREE-RANGING BOBCATS AND COUGARS IN THE UNITED

STATES: DISTRIBUTION, PREVALENCE, AND INTRASPECIFIC VARIATION

by

BARBARA C. SHOCK

(Under the Direction of Michael Yabsley)

ABSTRACT

Cytauxzoon felis, a tick-borne protozoal parasite (family Theileridae), is the causative

agent of cytauxzoonosis in domestic cats in the United States. The parasite was first identified in

domestic cats from Missouri, Texas, and Arkansas in the 1970’s and is now common in the

southeastern and Midwestern US. The bobcat (Lynx rufus) has been identified as the natural

reservoir of C. felis. The overall goal of this project was to better understand the natural history

of C. felis in bobcats and cougars (Puma concolor). Based on PCR testing of >700 wild felids,

infected bobcats were identified in numerous eastern and central states, with higher prevalence

rates being found in states where Amblyomma americanum is present in addition to

Dermancentor variabilis. Sequence analysis of internal transcribed spacer (ITS)-1 and ITS-2

regions revealed extensive genetic variation. Interestingly, one bobcat was infected with a

Babesia sp. which is the first report in a bobcat.

INDEX WORDS: Cytauxzoon felis, Babesia, Lynx rufus, Puma concolor, piroplasm

PIROPLASMS IN FREE-RANGING BOBCATS AND COUGARS IN THE UNITED

STATES: DISTRIBUTION, PREVALENCE, AND INTRASPECIFIC VARIATION

by

BARBARA C. SHOCK

B. S., West Virginia University, 2008

A Thesis Submitted to the Graduate Faculty of The University of Georgia in Partial Fulfillment

of the Requirements for the Degree

MASTER OF SCIENCE

ATHENS, GEORGIA

2010

© 2010

Barbara C. Shock

All Rights Reserved

PIROPLASMS IN FREE-RANGING BOBCATS AND COUGARS IN THE UNITED

STATES: DISTRIBUTION, PREVALENCE, AND INTRASPECIFIC VARIATION

by

BARBARA C. SHOCK

Major Professor: Michael J. Yabsley

Committee: Fred Quinn David S. Peterson

Electronic Version Approved:

Maureen Grasso Dean of the Graduate School The University of Georgia August 2010

iv

DEDICATION

I would like to dedicate this manuscript to my grandmothers, Betty Fern Weaver

and Jean Shock. Your support and love have been unending.

v

ACKNOWLEDGEMENTS

First I would like to thank Dr. Michael John Yabsley for being my major

professor. I have learned so much in the past two years from you. You are an engaging

and entertaining researcher, and I just hope that I can someday read a paper before you

have already. I look forward to working with you in the future.

My biggest thanks goes to my parents, Mark and Linda Shock for your support

and love. It’s always helpful to call home and have someone tell you they’re proud of

you. I would also like to especially thank my dad for helping me to collect samples for

this project and telling me about SCWDS. Although I have missed my family and friends

from home, they have always shown me the utmost support and love (this means you

Chelsie). I have to give a special thank you to Jenn and Todd Stueckle for first taking me

on in their lab as an undergrad and now for being my friends.

There are so many people at SCWDS that I am grateful to (and for), but I prefer to

thank you in person.

In addition, I’d like to thank numerous personnel from state agencies who

collected felid samples. The studies in this thesis were primarily funded by the Morris

Animal Foundation (DO8FE-003). Additional support was provided by the Federal Aid

to Wildlife Restoration Act (50 Stat. 917) and through sponsorship from fish and wildlife

agencies in Alabama, Arkansas, Florida, Georgia, Kansas, Kentucky, Louisiana,

Maryland, Mississippi, Missouri, North Carolina, Oklahoma, Puerto Rico, South

Carolina, Tennessee, Virginia, and West Virginia.

vi

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS .................................................................................................v

LIST OF TABLES ............................................................................................................ vii

LIST OF FIGURES ......................................................................................................... viii

CHAPTER

1 INTRODUCTION and LITERATURE REVIEW ...........................................1

Introduction .................................................................................................1

Literature Review: Tick-borne protozoa of felids ......................................2

Literature Cited .........................................................................................35

2 DISTRIBUTION AND PREVALENCE OF CYTAUXZOON FELIS IN

BOBCATS, LYNX RUFUS, FROM THIRTEEN STATES ........................49

3 EXTENSIVE GENETIC VARIABILITY OF CYTAUXZOON

FELIS FROM BOBCATS (LYNX RUFUS) AND COUGARS

(PUMA CONCOLOR) ..............................................................................65

4 NOVEL BABESIA IN A BOBCAT, GA ........................................................81

5 CONCLUSIONS.............................................................................................87

vii

LIST OF TABLES

Page

Table 2.1: PREVALENCE OF C. FELIS IN BOBCATS .................................................63

viii

LIST OF FIGURES

Page

Figure 1.1: PHYLOGENETIC RELATIONSHIPS BETWEEN GLOBAL SAMPLES

OF CYTAUXZOON FROM FELIDS ...................................................................................4

Figure 1.2: APPROXIMATE DISTRIBUTION OF DERMACENTOR VARIABILIS ......21

Figure 1.3: APPROXIMATE DISTRIBUTION OF AMBLYOMMA AMERICANUM .....21

Figure 2.1: DISTRIBUTION OF C. FELIS IN BOBCATS ..............................................64

1

CHAPTER 1

INTRODUCTION AND LITERATURE REVIEW

INTRODUCTION

Vector-borne pathogens are a significant cause of morbidity and mortality among people

and animals worldwide. Ticks are second only to mosquitoes as vectors of pathogens which

include a wide range of viruses, bacteria, and protozoa as well as some contribution to fungal

infection. Currently, there are three recognized genera of tick-borne protozoans, Theileria,

Cytauxzoon, and Babesia. The three are considered piroplasms because of their signet ring form

in erythrocytes. Babesia is the most numerous of the three and can be found all over the world in

mammals and birds. Theileria and Cytauxzoon are very closely related, and both also have a

global distribution. Infection with the piroplasms causes clinical signs which range from

subclinical parasitemia to mortality, depending on the type and strain of the parasite as well as

the host immune function.

Among felids, only Cytauxzoon and Babesia have been reported. Cytauxzoon felis and

Babesia felis are emerging significant pathogens in the United States and Africa, respectively.

Numerous other infections of felids with Cytauxzoon and Babesia spp. have been reported from

a variety of wild and exotic worldwide, but little work has been done to fully characterize these

parasites. In the United States, the reservoir species for C. felis is the North American bobcat

(Lynx rufus). Bobcats have been documented dying from infection with C. felis, but unlike the

high rates of mortality seen in domestic cats, most infected bobcats are subclinical carriers who

show little to no signs of infection. Due to the high prevalences of C. felis previously reported in

2

bobcats and the general lack of clinical signs, it is assumed that C. felis has been endemic in the

bobcat population for a long time. The overall goal of this project is to better understand the

natural history of C. felis in its wild felid reservoir. This goal had two focuses: quantifying the

distribution and prevalence of C. felis in wild felid populations; and characterizing the strains of

C. felis which are circulating in wild felids.

LITERATURE REVIEW: TICK-BORNE PROTOZOA OF FELIDS

Genera Cytauxzoon and Theileria

Cytauxzoon and Theileria species are two closely related tick-transmitted protozoan

parasites in the phylum Apicomplexa, class Aconoidasida, order Piroplasmorida, and family

Theilieridae (Kocan and Waldrup, 2001). Theileria are found in a wide range of hosts, primarily

ruminants, while Cytauxzoon is restricted to felids (Nijhof et al., 2005). Cytauxzoon are

distinguished from Theileria by the location of schizogony, with the cytauxzoons in felids

replicating in mononuclear phagocytes while theilerias in ungulates replicate in either

lymphocytes or mononuclear phagocytes (Ferris, 1997; Cowell et al., 1988; Preston et al., 1999;

Nijhof et al., 2005). Transmission of Cytauxzoon and Theileria occurs when sporozoites are

transferred to a vertebrate host during tick feeding. Experimental trials indicate transmission

occurs 3-5 days after tick-feeding commences (Hazen-Karr et al., 1987). Within the vertebrate

host, sporozoites first infect endothelial macrophages and undergo merogeny (Kocan et al.,

1992). Merozoites are released within 14 days, which then infect erythrocytes to form

intraerythrocytic piroplasms. These piroplasms are pleomorphic and can be round, oval, bipolar

or rod-shaped and range from 0.3 to 2.0 μm in diameter. Rarely, Maltese crosses and paired

3

piroplasms may be observed (Simpson et al., 1985; Kier et al., 1987; Kocan et al., 1992).

Transmission is completed when infected erythrocytes are ingested by a tick during feeding.

Although not well described, it is believed that sporogony occurs in the tick vector and that the

parasites migrate to the tick salivary glands prior to transmission. Experimental studies have

shown that the parasites are maintained transstadially (Blouin et al., 1984; Blouin et al., 1987;

Kocan et al., 1988).

The genus Cytauxzoon was first described in a gray duiker (Sylvicaprae grimmia) from

South Africa (Nietz and Thomas, 1948). Although numerous Theileria spp. had been described

before, the new genus was proposed because the parasite replicated in mononuclear phagocytes,

which had not been observed at that time in Theileria. The division of these parasites into

separate genera was a source of contention between Levine, who had classified the cytauxzoons

with the theilierias, and Brocklesby, who supported the separate genera based on the distinct cell

lineages used in schizogony (Brocklesby, 1979; Levine et al., 1980). Several other Cytauxzoon

spp. have been reported from African ungulates including: kudu, Tragelaphus strepsiceros;

eland, Taurotraugs oryx; and giraffe, Giraffa camelopardalis; however, these parasites are now

considered to be Theileria spp. (Neitz, 1964; Brocklesby, 1962; McCully et al., 1970; Nijhof et

al., 2005). Clinical morbidity and mortality ascribed to Cytauxzoon spp., but more likely

Theileria spp., has been reported in: tsessebe, Damaliscus lunatus; roan antelope, Hippotragus

equinus; sable antelope, H. niger; and suspected in an impala, Aepyceros melampus (Jardine,

1992; Bigalke, 1989; Wilson et al., 1974; Wilson et al., 1977; Carmichael and Hobday, 1975).

Currently Cytauxzoon spp. have been reported from domestic cats and wild felids in

seven countries including the United States, Brazil, Germany, France, Mongolia, Spain, and

Zimbabwe (Foggin and Roberts, 1982; Ketz-Riley et al., 2003; Mendes-de-Almeida et al., 2004;

4

Reichard et al., 2005; Luaces et al., 2005; Millan et al., 2007; Peixoto et al., 2007; Andre et al.,

2009; Criado-Fornelio et al., 2009). Molecular characterization suggest that multiple species of

Cytauxzoon infect felid species worldwide (Figure 1.1).

FIGURE 1.1. Phylogenetic relationships between global samples of Cytauxzoon from felids.

Sequences obtained from GenBank.

Genus Babesia

Babesia spp. are small piroplasms that have been detected in numerous mammalian hosts,

including several wild felid species. Babesia spp. are in the phylum Apicomplexa, class

Aconoidasida, order Piroplasmorida, and family Babesiidae. Similar to the Theileridae, Babesia

spp. are obligate intraerythrocytic protozoan parasites and are transmitted by Ixodid ticks. They

are distinguished from members of the Theileridae by the absence of tissue schizogenous stage.

Additionally Babesia spp. can also be transmitted transovarially in ticks while Theileridae are

Spain Iberian Lynx EF094469

Spain Iberian Lynx EF094468

France Domestic cat EU622908

Spain Iberian Lynx EF094470

Spain Domestic cat AY309956

Spain Iberian Lynx AY496273

Mongolia Pallas cat AF531418

Mongolia Pallas cat AY485691

Mongolia Pallas cat AY485690

Stabilate 153 (MO) L19080

Texas Domestic cat AY531524

Oklahoma Domestic cat AF399930

Theileria youngi Woodrat AF245279

Babesia sp. cougar DQ329138

100

100 98

10066

86

68

0.01

5

only transstadially transmitted between stages of ticks. Historically, Babesia are divided into two

morphologic groups, the “large” Babesia, which measure more than 2.5 μm, and the “small”

Babesia which measure 1.0 to 2.5 μm in size. Because of morphologic similarity among the large

and small piroplasms, genetic characterization is the best method to identify species (Kocan and

Waldrup, 2001).

A number of Babesia spp. have been recognized in felids from around the world. Large

Babesia include B. herpailuri and B. pantherae from wild felids in Africa and small Babesia

include B. felis and B. leo from domestic cats and other felids in Africa, B. cati in domestic cats

from India, B. canis canis from domestic cats in Spain, B. canis presentii from domestic cats in

Israel; a Babesia sp. from domestic cats in Portugal (called Theileria annae), and a Babesia sp.

from Florida panthers in the United States (Jacobson et al., 2000; Criado-Fornelio et al., 2003;

Penzhorn et al., 2004; Criado-Fornelio et al., 2004; Baneth et al., 2004; Yabsley et al., 2006).

Domestic feline babesiosis is generally a mild chronic disease, although the pathogenicity

depends on the species or sometimes the strain of the parasite (Penzhorn, 2006). The most

common clinical signs include anorexia and lethargy. When disease is present, Babesia spp.

generally cause hemolytic anemia, but most felids do not develop clinical disease unless

presented with a secondary or immunocompromising infection or stressor (Shoeman et al.,

2001). However, exceptions occur as B. felis is a significant pathogen of cats in South Africa

(Penzhorn et al., 2004).

6

Cytauxzoon spp. in felid species from the United States

Domestic cats

Cytauxzoonosis was first recognized in domestic cats (Felis domesticus) in Missouri,

Arkansas, and Texas during the early 1970’s by the presence of piroplasm-infected erythrocytes

and microscopic lesions (schizont-laden cells lining the walls of blood vessels) as well as

piroplasm-infected erythrocytes (Bendele et al., 1976; Wagner, 1976; Wightman et al., 1977). A

similar syndrome had been previously recognized in African ungulates for many years (Neitz

and Thomas, 1948; Neitz, 1957; Martin and Brocklesby, 1960; McCully et al., 1970; Wilson et

al., 1974); thus these feline infections represented the first reports of a Cytauxzoon-like illness in

the United States as well as the first report in a felid. Based on previous research involving

Theileria and Cytauxzoon, the parasite was presumed to be tick-transmitted. The parasite,

Cytauxzoon felis, has now been described in domestic cats from Alabama, Arkansas, Florida,

Georgia, Kansas, Kentucky, Louisiana, Mississippi, Missouri, North Carolina, Oklahoma, South

Carolina, Tennessee, Texas, and Virginia (Bendele et al., 1976; Wagner, 1976, Wightman et al.,

1977; Ferris, 1979; Glenn and Stair, 1982; Hauck, 1982; Kocan and Kocan, 1991; Meier and

Moore, 2000; Birkenheuer et al., 2006; Haber et al., 2007).

Bobcats

Piroplasms morphologically consistent with C. felis was described in bobcats as early as

1930 (Wenyon and Hamerton, 1930) and were again observed in two free-ranging bobcats when

C. felis was first recognized in domestic cats (Wagner, 1976). To better characterize this

piroplasms, researchers at the University of Missouri, College of Veterinary Medicine tested the

ability of C. felis to infect 30 different domestic, laboratory, and wildlife species (Kier et al.,

7

1982a). Inoculation of blood, spleen and lymph node or tissue homogenate from experimentally

infected domestic cats into laboratory mice (Mus musculus, ICR), immunosuppressed nude

laboratory mice (M. musculus, BALB/c-nu), rats (Rattus norvegicus), gerbils (Meriones

unguiculatus), hamsters (Mesocricetus auratus), guinea pigs (Cavia porcellus), a chinchilla

(Chinchilla laniger), rabbits (Oryctolaugs cuniculus), squirrel monkeys (Saimiri sciureus), a dog

(Canis familiaris), a cow (Bos Taurus), a goat (Capra hircus), a sheep (Ovis aries), a pig (Sus

scrofa), coyotes (Canis latrans), a fox (Vulpes fulva), skunks (Mephitis mephitis), raccoons

(Procyon lotor), a woodchuck (Marmota monax), a marmot (Marmota flaviventris), opossums

(Didelphis marsupialis), ground squirrels (Citellus tridecemlineatus), a grey squirrel (Sciurus

carolinensis), a vole (Microtus ochrogaster), white-footed mice (Peromyscus maniculatus), bats

(Myotis lucifugus), cottontail rabbit (Sylvilagus floridanus), a deer (Odocoileus virginianus), an

ocelot (Felis pardalis), a mountain lion (P. concolor), and bobcats resulted in infections only in

domestic sheep and the two bobcats. It is unknown why the domestic cats, ocelot, and mountain

lion failed to develop infections. The inoculated sheep developed a low but persistent

parasitemia, but no clinical signs of disease. However, domestic cats inoculated with tissue

homogenates from the sheep did not develop cytauxzoonosis.

Numerous studies have established that bobcats are experimentally and naturally

susceptible to infection with C. felis. In addition, variability in disease progression in

experientially infected bobcats has been observed that may be related to infectious dose, route, or

stage or subspecies of bobcat, a combination of both, or some other factor (Kier et al., 1982b;

Kier et al., 1983). The first experimental inoculation trial gave very different results for two

different bobcats (Kier et al., 1982a; Kier et al., 1982b). One bobcat from Florida (here termed

Lynx rufus floridanus), developed clinical signs of cytauxzoonosis including anorexia,

8

depression, and parasitemia, and died two weeks post-inoculation. Histological examination of

tissues revealed large numbers of schizonts occluding vessels. The second bobcat (here termed

Lynx rufus rufus) developed a subclinical parasitemia. The eastern bobcat, which survived the

initial infection with C. felis, was inoculated four subsequent times, none of which resulted in

clinical signs. However, the bobcat did develop transiently elevated infected red blood cell

(IRBC) counts and after splenectomy and steroid treatment, the bobcat developed moderate

anemia, reticulocytosis, and elevated IRBC counts. This bobcat maintained a subclinical

parasitemia for 1,372 days (approximately 4 years) before dying of congestive heart failure.

Upon necropsy, no schizogenous states of C. felis were observed and the final parasitemia was

8.5% (Kier et al., 1982b). Domestic cats inoculated with blood from these two bobcats also

presented with variable disease: cats inoculated with blood from the surviving eastern bobcat

survived while cats inoculated with blood from the Florida bobcat died. This study had three

important findings: 1) bobcats can develop disease, 2) bobcats can have long-term parasitemias,

and 3) domestic cats can survive experimental infection.

Simultaneously, a survey of blood smears from bobcats from Oklahoma showed that 13

of 21 (62%) were positive for Cytauxzoon-like piroplasms (Glenn et al., 1982). The average

parasitema observed was 1-3%, but ranged from 0.5% to 5%. Of the 13 positive bobcats, one

exhibited anemia, a classic sign of cytauxzoonosis in the domestic cat. A subsequent study

collected blood from four wild-caught bobcats which had a naturally occurring infection with an

intraerythrocytic piroplasm and showed that the parasites caused fatal cytauxzoonosis in a single

domestic cat (Glenn et al., 1983). Three other inoculated cats developed subclinical infections

(Glenn et al., 1983). A single cat that was subsequently inoculated with a domestic cat-origin

virulent inoculum of C. felis developed fatal cytauxzoonosis within 14 days. Two of the original

9

wild-caught bobcats were also inoculated with the same strain of C. felis as the domestic cat, but

they did not develop clinical disease. Histologically, schizogenous forms of C. felis were not

observed in any tissues from the bobcats or from the two subclinically infected domestic cats.

Examination of 10 naturally-infected bobcats revealed that schizonts of C. felis were

absent from the liver, lungs, spleen, and lymph nodes. Spleen homogenates from four of the

bobcats were inoculated into domestic cats. Schizonts of C. felis were not seen in any tissues of

the naturally infected free-ranging bobcats. Additionally, the domestic cats inoculated with the

spleen homogenate did not develop clinical signs of cytauxzoonosis, but did develop subclinical

parasitemias. Later the same group studied the clinical progression of C. felis in bobcats

following exposure to infected D. variabilis (Blouin et al., 1987).

In the second part of the study (Blouin et al., 1987), the researchers splenectomized a

naturally infected bobcat which had a previous parasitemia of 1%. After the splenectomy, a

spleen homogenate was inoculated into a domestic cat. D. variabilis nymphs were allowed to

feed on this bobcat. Two bobcats (no. 2, 3) which had been determined to be uninfected with C.

felis were parasitized by the infected ticks. Prescapular lymph nodes were removed from the two

bobcats at 11 days post-exposure for impression smears and inoculation into domestic cats. A

second prescapular lymph node was removed from bobcat 3 after 30 days post-exposure, and

smears and inoculation were performed. The first, splenectomized, bobcat displayed no clinical

signs of C. felis and schizogenous stages of C. felis were not observed in a histological

examination of the spleen. Schizogenous stages were observed in reticuloendothelial

macrophages in the 11 day post-tick attachment lymph node impression smears of bobcats 2 and

3 and the domestic cats which were inoculate with these tissue homogenates died of acute

cytauxzoonosis 11 and 14 days post-inoculation respectively. Interestingly, bobcat 2 developed

10

clinical signs of cytauxzoonosis 19 days post-tick attachment and died. Gross and microscopic

lesions revealed classical signs of cytauxzoonosis. The lymph node impression smear from

bobcat 3 which was made 30-days post-tick exposure revealed no schizogenous stages of C. felis

and the domestic cat inoculated with this tissue homogenate developed a parasitemia of 2-3% but

showed no clinical signs of C. felis. Gross and microscopic evaluation of bobcat 3 after

euthanasia 60 days post-tick attachment revealed no schizogenous stages of C. felis and only a

8% erythrocytic parasitemia. This paper reveals that development of the schizogenous phase in

bobcats can be initiated by sporozoites transmission from the tick vector. It is also evidence that,

in general, schizogenous development of C. felis in bobcats is limited and the erythrocytic stage

is maintained and dominant. Due to the death of one bobcat (no. 2) in this study from tick-

transmitted C. felis, and a previous experimental death of a bobcat (Kier et al., 1982a; Kier et al.,

1982b), it can be assumed that while most bobcats in the wild probably have limited

schizogenous stages, some may die of naturally acquired C. felis infections.

Although a few experimental studies (Kier et al., 1982b; Blouin et al., 1984) indicated

that bobcats could develop clinical disease, evidence for natural mortality is limited. Only a

single case has been reported (Nietfeld and Pollock, 2002). In 2000, a moribund free-ranging

bobcat kitten collected from an open field near Wamego, KS had severe anemia and respiratory

difficulty (Nietfeld and Pollock, 2002). Gross examination revealed multifocal petechiae,

splenomegaly, and pericardial effusion. Histological examination revealed subacute pulmonary

thrombosis, mild vasculitis in the brain, and schizont-filled macrophages occluding the blood

vessels of all examined tissues.

The first study to examine the prevalence of C. felis in bobcat populations based on PCR

testing was Birkenheuer et al. (2008) who tested bobcats from North Carolina and Pennsylvania.

11

Currently, C. felis has not been reported from domestic cats in Pennsylvania, but has been

reported numerous times from domestic cats in North Carolina (Birkenheuer et al., 2006a).

Based on PCR, samples were collected from legally-trapped bobcats in 33% of 32 bobcats from

North Carolina and 7% of 70 bobcats from Pennsylvania were positive for C. felis. This

represented the first report of C. felis from any felid in Pennsylvania and the authors alerted

veterinary practitioners to the possibility of C. felis infection in domestic cats in that state.

The high prevalence and low pathogenicity of C. felis for bobcats suggests that C. felis

has been endemic in the populations for some time. As no Babesia or Theilieria spp. have yet

been described in bobcats, it can be assumed the piroplasm described in a bobcat in 1930 by

Wenyon and Hamerton was in fact C. felis and that the infection has been endemic in the

population since at least the early 1900s. As the distribution of A. americanum and D. variabilis

changes over time, it is probable that naïve populations of bobcats will become exposed to this

parasite. As Birkenheuer et al. (2008) stressed, further studies are warranted to better understand

the natural history and epidemiology of C. felis in bobcat populations.

Florida panthers

The first report of C. felis in a Florida panther (Puma concolor coryi or Felis concolor

coryi) was in 1991. The detection was accidental and occurred when researchers with the Florida

Game and Fresh Water Fish Commission and Cornell University unknowingly infected a

domestic cat with C. felis during a study to determine if a 3 yr-old female panther had feline

immunodeficiency virus (FIV) (Butt et al., 1991). The researchers intraperitoneally inoculated a

FIV antigen and antibody-negative adult domestic cat with 1.5x106 mononuclear cells from the

panther and 11 days later the cat became depressed and febrile. Although supportive care was

12

administered, the cat died the next day and upon necropsy, gross lesions such as petechial

hemorrhages and splenic enlargement were noted. Microscopic examination of tissues revealed

schizont-filled mononuclear cells which completely occluding vessels. Examination of blood

smears indicated that the cat had a 10% parasitemia (Butt et al., 1991). Retrospective analysis of

blood smears from the infected Florida panther revealed she was infected; however, the panther

had received two blood transfusions from other P. concolor donors, so it was unclear if this

infection was natural or iatrogenic.

A subsequent study was conducted to determine the prevalence of C. felis in the Florida

panthers and introduced Texas cougars (Puma concolor stanleyana) (Rotstein et al., 1999). The

Texas cougars included in the study were translocated into Florida to introduce additional

genetic variability and to increase the population size, which at that point was estimated to be

~50 panthers (Maehr, 1997). A retrospective analysis (1983-1997) of blood smears revealed that

39% of the Texas cougars (11/28) and 35% of the Florida panthers (22/63) were infected with C.

felis. Interestingly, a 7 day-old kitten was positive for C. felis. No difference in prevalence was

noted between the sexes. Infected Florida panthers had significantly lower mean cell hemoglobin

and monocytes counts and significantly higher neutrophil and eosinophil counts compared with

infected Texas cougars. However, all the values were within the normal range of expected values

so the authors concluded that there was no biological significance associated with the differences

(Rostein et al., 1999). These data suggested that the Florida panther may be a reservoir for C.

felis.

To better understand the role of Florida panthers as a potential reservoir, a PCR-based

survey was conducted (Yabsley et al., 2006). Amplification of a conserved segment of the 18S

rRNA gene revealed that 39 of 41 panthers were infected, which was significantly more than

13

indicated by blood smear analysis (only 15 infected). Surprisingly, after sequence analysis, the

majority of the panthers (95%, 32 of 39 infected panthers) were infected with a novel Babesia sp.

and only seven were infected with C. felis (Yabsley et al., 2006). Two of the cougars were co-

infected with C. felis and the Babesia sp. This represented the first report of a Babesia sp. in a

felid from North America.

Because two morphologically similar piroplasms had been reported in the Florida

panther, the conclusions of Rotstein et al. (1991) were re-examined. Although, fatal

cytauxzoonosis has not been described in a Florida panther, Harvey et al. (2007) described three

cases of cougars that had acute C. felis infections that resulted in mild clinical signs and mild

hematologic and biochemical abnormalities. Of the three case reports in Harvey et al. (2007),

one cougar demonstrated clinical signs after infection (year 1989) including anorexia,

depression, and dehydration. This cougar was also infested with A. americanum, a confirmed

vector of the C. felis (Reichard et al., 2009). Sequential testing of blood indicated that the cougar

remained infected for at least one year while in the Florida facility and was still infected in 2005

when it was euthanized due to diabetes at her new home in Savannah, Georgia. It is unknown if

she maintained the same subclinical C. felis infection for at >16 years or if she acquired a new

infection while in Georgia. The other two cases were western cougars that were part of the

genetic restoration study for the Florida panthers. These cougars were housed in northern Florida

in 1995 before their release into southern Florida and were negative for C. felis (and Babesia sp.)

during the initial 3-week quarantine period. After their third week at the facility, the cougars

developed anemia and had become infected with C. felis. Neither cougar showed clinical signs

and both were released in southern Florida. The cougars died in 1999 and 2001 of illnesses

unrelated to cytauxzoonosis, although both were PCR positive for C. felis at the time of death.

14

These data indicated that during acute infections with C. felis, cougars can develop mild

hemolytic anemia as well as liver injury, but they to survive and develop subclinical infections.

Exotic felids

Acute fatal cytauxzoonosis has been reported in a white tiger (P. tigris) (Garner et al.,

1996) that was housed at the same facility which housed many of the infected cougars from

previous studies (Butt et al., 1991; Rotstein et al.,1991; Harvey et al., 2007). The 7-year-old

female had a two-day history of anorexia and lethargy and on the third day of illness, two female

A. americanum were found and removed. She subsequently developed icterus, a low hematocrit

(26%), and was mildly dehydrated. Two days later, she was recumbent and developed petechiae

and profuse bleeding at puncture sites. She also had a parasitemia of 5% and a mild

nonregenerative anemia, moderate leucopenia, neutropenia, lymphopenia, and severe

thrombocytopenia. Histologically, C. felis was present in mononuclear phagocytes which were

found in large quantities occluding capillaries as well as small arteries and veins (Garner et al.,

1996).

Infection dynamics, clinical signs, and pathology of C. felis

The classic clinical signs observed from C. felis in domestic cats begin with anemia and

depression and are quickly followed by fever, dehydration, icterus, splenomegaly, and

hepatomegaly. The pathognomonic signs of C. felis infection are erythrocyte hemolysis and

occlusion of the lumen of blood vessels by large schizont-laden mononuclear phagocytes in the

lungs, liver, lymph nodes, and spleen (Simpson et al., 1985; Kier et al., 1987; Kocan and Kocan,

1991; Kocan et al., 1992).

15

A large group of researchers from the University of Missouri, which had first identified

the illness in domestic cats, and the Plum Island Animal Disease Center (PIADC) investigated

experimental routes of infection in the domestic cat (Wagner et al., 1980). Experimental

infection trials with 131 domestic cats were conducted with parenteral administration of either

fresh or liquid nitrogen-frozen blood or tissue homogenates that were collected from domestic

cats with acute cytauxzoonosis. Two cats were splenectomized to determine if splenectomization

enhanced the infection. Two uninoculated cats were held with the inoculated cats until they died

of cytauxzoonosis and then a further 60 days to determine if the parasite could be transmitted

directly from cat to cat. Additionally, one cat was administered tissue homogenate via gastric

lavage, while the same suspension was intraperitoneally inoculated into another cat to determine

if the parasite could be obtained by ingestion. This study determined that domestic cats do not

acquire C. felis via contact or ingestion of infectious tissues. This study also determined that the

minimum infectious dose of C. felis for a domestic cat is a 0.25mL subcutaneous inoculation of a

1:10 dilution of frozen standardized spleen inoculum. The standard progression to acute

cytauxzoonosis was between 17 and 20 days, with an average of 18.4 days. Of the 131 cats

included in this study, 54% died of clinical cytauxzoonosis and 46% were euthanized due to

acute cytauxzoonosis. The most frequent clinical signs were pyrexia, depression, anorexia and

dehydration initially followed by fever, lethargy, severe dehydration, icterus then coma and

death (Wagner et al., 1980).

Concurrent with the Wagner et al., (1980) study, researchers at the PIADC

experimentally infected >500 domestic cats and numerous domestic livestock species with C.

felis to elucidate the relationship between this piroplasm and the organisms observed in African

ungulates and to address the potential threat of this parasite to livestock in the United States

16

(Ferris, 1979). The study was discontinued after C. felis was proven to not be a threat to domestic

livestock; however, these data significantly added to the knowledge of C. felis in the United

States. Importantly, one of over 500 domestic cats that were infected in this larger study survived

which was the first report of a domestic cat surviving infection with C. felis. Additionally,

researchers concluded that C. felis can be experimentally transmitted by inoculation of as little as

0.2 ml of blood, splenic, hepatic, pulmonary and lymph node homogenates by any route of

injection (eg. intravenous, intraperitoneal, subcutaneous, intradermal, etc). They confirmed that

initial clinical signs, depression and anorexia, were typically seen 5-7 days post infection and

that fever slowly rose to 40-41.1 C and stays at that level an average of 3-5 days before dropping.

Before death, pyrexia, anemia, icterus and dehydration as well as dyspnea were observed and

death usually occurred one to two weeks after development of clinical signs. After necropsy,

gross examination revealed splenomegaly and petechial hemorrhages on lymph nodes and lungs.

Microscopic evaluation revealed schizogenous stages of the parasite in reticuloendothelial

histocytic cells which occluded blood vessels. The lungs were the organ most heavily affected,

followed by the spleen, and then the liver and lymph nodes. The highest parasitemia observed in

this study was 4% with an average of 1%, but researchers in Missouri had reported a parasitemia

as high as 25% (Ferris, 1979; Wagner et al., 1980).

The pathology of experimental infections of domestic cats with C. felis was more

completely described by Kier et al. (1987). Experimentally infected cats were sacrificed from

day 1 to day 19 post-inoculation (PI) for histological evaluation. By day 16 PI cats were

exhibiting classic clinical signs of infection. Parasitemias were first noted 10 days PI. The first

schizogenous stage of C. felis was observed at day 12 PI and there was a significant correlation

between levels of parasitemia with temperature rise, presence of tissue stages, and decreased

17

white blood cell counts. Recently, a retrospective study of the pulmonary histopathology of C.

felis infection was conducted on 148 domestic cats from January 1995 to June 2005 in Oklahoma

(Snider et al., 2010). Interstitial pneumonia was found to be moderate in most cases, numbers of

alveolar macrophages and intra-alveolar hemorrhages were low, and in many cases, infiltrations

of neutrophils were noted. Similar to previous studies, extensive vascular occlusion, one of the

hallmark signs of acute cytauxzoonosis, was noted (Snider et al., 2010).

Both in situ hybridization and immunohistochemical techniques have been used to study

the pathogenesis of cytauxzoonosis in domestic cats (Susta et al., 2009). Using a riboprobe

targeting the 18S rRNA region of Babesia microti, C. felis-infected cells were most often

observed in the pulmonary intravascular macrophages and alveolar macrophages and

macrophages in the spleen, but ISH-positive cells were also seen in the kidneys, heart, and brain.

Using immunohistochemistry with a monoclonal antibody (Mac387), C. felis-infected cells were

found to be negative for calprotein which indicates a decrease in diapedesis which would provide

more circulating parasites available to the tick. Immunohistochemistry of two proliferation

markers, the proliferating cell nuclear antigen and p53, also showed that infected cells replicate

more frequently. While C. felis is in the cytoplasm, it blocks the translocation of the p53

proapoptotic protein, to the nucleus which would prevent the cell from undergoing apoptosis

(Susta et al., 2009).

A single case of abortion and death due to cytauxzoonosis in a domestic cat from Georgia

has been reported (Weisman et al., 2007). The 1-year old cat was in her 5th or 6th week of

gestation when she aborted and died. Fetal tissue, including skeletal muscle, developing bone

and bone marrow, and placenta were negative for piroplasms and schizogenous stages of C. felis.

Although it is unknown if C. felis can be transmitted transplacentally, there is evidence for

18

transplacental transmission of T. equi and T. sergenti in horses and cows, respectively (Baek et

al., 2003; Phipps and Otter, 2004; Allsopp et al., 2007).

Diagnostic testing for C. felis

Previously, the standard method of diagnosis for C. felis infection was blood smear

analysis to detect the piroplasm infected red blood cells (Glenn et al., 1982; Kier et al., 1982a;

Ferris, 1979). Recently, molecular methods have been used to detect infection with greater

sensitivity. Polymerase chain reaction (PCR) can be used to detect low numbers of parasites and

to characterize the piroplasms found (Birkenheuer et al., 2006; Brown et al., 2008). Serologic

tests for C. felis have been investigated, but domestic cats typically have high numbers of

intraerthrocytic forms that are easily observed during acute infections. With the increased

recognition of chronically infected cats, however, serologic testing might be useful for

population-based studies (Shindel et al., 1978; Cowell et al., 1988).

Treatment and survival in C. felis

Numerous treatment regimes have been tested including parvaquone, imidocarb,

dimiazene aceturate, buparvaquone; however, imidocarb is currently the drug of choice (Green et

al., 1999). Although several cats survived infection during these clinical trials (4 of hundreds), no

treatment, even imidocarb, has been considered 100% effective and consistent in managing the

protozoan infection. In 1987, Uilenberg et al. inoculated a domestic cat with C. felis and treated

it with parvaquone. This cat was subsequently immune to the same inoculum. Another study

used a combination of parvaquone and buparvaquone in 17 cats, and although one infected cat

survived, another cat not treated also survived (Motzel and Wagner, 1990). Importantly, data

19

indicate that cats that are treated can remain parasitemic which might allow them to continue the

life cycle by infecting ticks (e.g. three cats in Arkansas that were treated with imidocarb

dipropionate remained parasitemic for 7 months, 15 months, and 29 months) (Brown et al.,

2008).

Historically, this disease was considered by be nearly uniformly fatal for domestic cats,

but over the years there have been indications (during experiment studies or field-based studies)

that some cats could survive and develop subclinical chronic infections (Ferris, 1979; Meinkoth

et al, 2000; Brown et al., 2008; Haber et al., 2007). Increased recognition of subclinical

infections suggests that either cats are adapting to the parasite, less virulent strains of the parasite

are beginning to emerge in domestic cats, or better diagnostic assays are detecting these chronic

asymptomatic cases. From 1997 to 1998, Meinkoth et al. (2000) reported 18 cats from Arkansas

and Oklahoma which had survived natural infections with C. felis. Four of the cases were

asymptomatic and were identified only after one of their housemates showed clinical signs of

cytauxzoonosis; these cats had shown no clinical signs. Only one of the cats was treated with

imidocarb and all the cats were still parasitemic after 154 days. A single cat tested after 6 yrs was

still infected (from Walker and Cowell, 1995). Three other studies have reported a number of

cats that either survived infection or were chronic carriers including two of 34 cats in North

Carolina, two of 961 cats from Florida, three cats in Arkansas, and one of 75 cats from

Tennessee (Birkenheuer et al., 2006; Haber et al., 2007; Brown et al. 2008). Importantly, these

data suggest that domestic cats can develop long-term parasitemias which could allow them to

serve as reservoirs for the parasite.

20

Vectors of Cytauxzoon felis

In experimental studies, C. felis has been transmitted by two Ixodid tick species,

Dermacentor variabilis, the American dog tick, and Amblyomma americanum, the lone star tick.

D. variabilis is a moderate-sized tick (2-6 mm long) that has a three-host lifecycle involving

small mammals for the larval and nymph stages and large mammals for the adult stage (Allan,

2001). D. variabilis has a distribution involving the entire Eastern United States, the Pacific

coast, and parts of Idaho and Montana (Allan, 2001) and is primarily found in areas with forest

undergrowth (Allan, 2001; Figure 1.2). Ticks of the genus Amblyomma are larger ticks (4-8mm),

that tend to be catholic in their feeding habits, with three all stages feeding on a variety of

different hosts. In general, the genus is restricted to areas of warmer temperature and high

humidity for development of each life stage (Semtner et al., 1973; Koch and Dunn, 1980b). The

distribution of A. americanum ranges primarily in the Southeastern and south-central United

States, although new reports indicate range expansion (Allan, 2001, Figure 1.3) into coastal

regions of Maine and other Northeastern states (Keirans and Lacombe, 1998). The preferred

habitat of A. americanum is subclimax forests (Sonenshine and Levy, 1971).

21

FIGURE 1.2. Approximate distribution of Dermacentor variabilis (CDC, 2009)

FIGURE 1.3. Approximate distribution of Amblyomma americanum (CDC, 2009)

C. felis was first experimentally transmitted from a bobcat to a domestic cat by D.

variabilis (Blouin et al., 1984). Lab-raised nymphs were fed to repletion on a splenectomized

22

bobcat with a parasitemia of 40%, allowed to molt to the adult stage, and then were fed on

splenectomized domestic cats. Both domestic cats died of acute cytauxzoonosis at 13 and 17

days post-tick engorgement. Tissue impression smears displayed the schizogenous stage of C.

felis and the domestic cats displayed clinical and microscopic evidence of infection with C. felis.

Inoculation of a domestic cat with parasitemic blood from the same bobcat resulted in a

subclinical infection which was maintained for 6 months. This study was the first to show that D.

variabilis transstadially maintains C. felis and can transmit the parasite to felids. The study also

showed that clinical outcome in domestic cats may be related to the route of infection because

blood-inoculation (as was seen previously) caused subclinical infection whereas tick-

transmission produced acute cytauxzoonosis (Blouin et al., 1984).

D. variabilis also successfully transmitted C. felis infection from a splenectomized

parasitemic bobcat to two un-splenectomized bobcats which were previously determined to be

uninfected with C. felis by both blood smears and inoculation of whole blood into domestic cats

(Blouin et al., 1987). One of the bobcats died 19 days post tick-attachment of clinical

cytauxzoonosis which was the first indication that bobcats could develop clinical disease.

The first survey of ticks for C. felis infection was conducted by Bondy et al. (2005) who

screened 1, 362 ticks (Rhipicephalus sanguineus, D. variabilis, A. americanum) from Missouri.

Unfortunately, all the ticks for this study were collected from domestic dogs and cats which

complicated interpretation of the results. Of all the samples included in the study, only three

(0.93%) A. americanum nymphs were PCR positive for C. felis; however, all three were

collected from a domestic cat which was confirmed to be infected with C. felis. Although this

study suggested a role of A. americanum in the transmission of C. felis to domestic cats, there

were several caveats which prevented the A. americanum from being confirmed as a vector.

23

First, since the ticks were removed from a C. felis positive individual, it is impossible to

determine if the felid acquired the infection from the A. americanum, if the ticks were positive

because they were engorged with C. felis-infected feline blood, or if the vectors were positive

because they had acquired the parasite from a previously infected felid. Secondly, clinical signs

of infection usually do not develop for at least two weeks post-tick engorgement so the

likelihood of one of these nymphs being the vector is very low (Bondy et al., 2005; Blouin et al.,

1987; Blouin et al., 1984).

Nevertheless, A. americanum was suspected to be a vector because the ubiquitous nature

of the tick in regions where C. felis had been identified in domestic cats (Reichard et al., 2008)

and epidemiologically, peaks of cytauxzoonosis in domestic cats in May and September

correlated with natural peaks in A. americanum activity (Reichard et al., 2008). A. americanum

was confirmed as a competent vector when Reichard et al., (2009) conducted a transmission trial

using lab-raised A. americanum, D. variabilis, R. sanguineus, and Ixodes scapularis nymphs that

had fed on a subclinically C. felis-infected cat. Only the cat infested with adult A. americanum

exhibited clinical signs of cytauxzoonosis (11 dpi), while none of the other cats infested with

adult D. variabilis, R. sanguineus, and I. scapularis became infected. Similarly, in a subsequent

study D. variabilis failed to transmit C. felis (Edwards et al., 2010). Although D. variabilis did

not transmit C. felis in these two studies, there are a number of differences between these studies.

The parasitemia of the initial felids used in the studies (40% in Blouin et al., (1984) vs. 0.015%

in Reichard et al., (2008)) and the immune status of the subject felids (splenectomized vs. un-

splenectomized).

After A. americanum was confirmed to be a vector, a field-based study of wild-caught

questing ticks from natural habitats surrounding Stillwater, Oklahoma was conducted. C. felis

24

was detected in A. americanum (MIRs of 0.5% (1 of 178) for adult males, 1.5% (3 of 197) for

adult females, and 0.8% (3 of 393) for nymphs (Edwards et al., 2010). All 160 D. variabilis were

negative.

Currently, both D. variabilis and A. americanum are considered competent vectors of the

parasite (Blouin et al. 1984; Blouin et al., 1987; Reichard et al., 2008); however, very little is

understood about the importance of each vector species in the overall ecology of C. felis. Based

on A. americanum densities in areas where cytauxzoonosis is common and the epidemiologic

association with A. americanum activity, A. americanum likely represents the primary vector of

C. felis.

Seasonality and risk factors for C. felis

Because cytauxzoonosis is a tick-borne disease, a marked seasonality in diagnosed cases

has been noted. A retrospective analysis of cases submitted to the Oklahoma Animal Disease

Diagnostic Laboratory (n=180) during 1995-2006 and the Boren Veterinary Medical Teaching

Hospital (n=52) during 1998-2006 found a bimodal pattern with a large peak in the spring and

early summer months of April, May and June, and a small peak in the autumn months of August

and September (Reichard et al., 2009). These peaks correspond with the activity of the tick

vectors, especially A. americanum.

The authors were able to identify the geographic coordinates and landscape

characteristics for 41 of the cases and 68.3% were reported to occur in low density residential

areas. More cases (19.5%) occurred in urban edge habitat than expected at random. Significantly

more cases of cytauxzoonosis were associated with wooded cover and proximity to natural or

unmanaged areas. This study is a confirmation of observations clinicians and researchers had

25

been making for years about the risk factors associated with cytauxzoonosis in domestic cats.

Basically, cats which reside in habitats that support tick vectors, bobcats or both are more likely

to become infected with C. felis. Importantly, risk of C. felis infection is greatly reduced (nearly

preventable) by limiting tick exposure by keeping cats indoors.

Genetic characterization of Cytauxzoon felis from domestic cats

There have been at least three reasons postulated for the increased recognition of

chronically infected asymptomatic cats: 1) better treatment strategies, 2) better diagnostics, or 3)

variable strains of C. felis that differ in their virulence for domestic cats. Although many

treatment strategies have been attempted in reducing the parasitemia in domestic cats (Green et

al., 1999; Motzel and Wagner, 1995), none have been consistently effective and recently there

have been increasing reports of natural subclinical chronically-infected domestic cats

(Birkenheuer et al., 2006; Brown et al., 2008). Although diagnostics, particularly molecular

techniques, have improved, historically experimental studies have suggested that different

strains of C. felis had variable pathogenicity for cats as some strains induced clinical disease

while others induced subclinical chronic infections (Kier et al., 1982b).

If genetic markers could be identified to identify clinically different strains, then the

clinical outcome and treatment protocols might be more easily predicted (Brown et al., 2009a).

One of the most commonly used markers for this type of analysis is the noncoding first and

second internal transcribed spacer regions of the ribosomal RNA operon (ITS-1 and ITS-2).

These targets are more likely to have genetic variability compared with the conserved regions of

this operon (e.g., 18S, 5.8S, and 28S rRNA) (Hills and Dixon, 1991). The ITS-1 and ITS-2

regions have been useful in examining variability among a related hemoparasite, Babesia canis,

26

as well as two other apicomplexans, Cyclospora cayetanensis and Eimeria spp. (Brown et al.,

2009a; Barta et al., 1998; Olivier et al., 2001; Zahler et al., 1998).

From 2005 to 2007, samples of C. felis from domestic cats from Arkansas and Georgia

were genetically characterized and analyzed by clinical outcome (Brown et al., 2009a).

Unambiguous ITS-1 and ITS-2 sequences of C. felis were obtained for 88 of 112 C. felis

samples. These sequences were divided into three distinct genotypes (ITSA, ITSB, ITSC) each

of which were detected in at least five cats. Additionally, eight other unique sequences were

noted (Brown et al., 2009a). Within the ITS-1 region (~458bp), variability was due to eight

single nucleotide polymorphisms (SNPs) and one single nucleotide insertion and within the ITS-

2 region (~265bp), variability was due to four SNPs and one 40 base pair insertion. ITSA was

the most commonly identified genotype (54.5% of samples) followed by ITSB (23.9%), and

ITSC (5.7%). Of cats infected with the ITSA genotype, a significant number (38/48, 79.2%)

survived infection compared to those infected with the ITSB genotype (4/21, 19.0%) and the

ITSC genotype (0/5, 0%). These data suggested that specific ITS genotypes may be associated

with clinical outcome in domestic cats (Brown et al., 2009a).

In addition, most of the cats that survived C. felis infection were from the Midwest and

the ITSA genotype was identified in 84.2% of Arkansas samples but from only one sample

collected from Georgia. Conversely, ITSB was identified in 67.7% of the samples from Georgia,

but no samples from Arkansas. ITSC was only detected in samples from Georgia (5/57, 8.8%).

Thus, these data also indicate that certain strains of C. felis circulate in different geographic

regions.

Recently, a retrospective study (1995-2007) of 98 C. felis histology specimens from

Georgia found similar results (Brown et al., 2009b). Although 85 of the 98 samples yielded

27

unambiguious ITS-2 sequences, only 21 samples yielded near complete ITS-1 region sequences.

To increase numbers, the ITS-1 sequence analysis was restricted to 290 base pairs, which

allowed a total of 48 samples to be included in the study. Three identified genotypes (ITSa,

ITSb, and ITSc) were detected as well as 8 additional unique sequences. Twenty-seven of the

samples were ITSa, 8 were ITSb, and 3 were ITSc. One of the eight new sequences (described as

ITSd) was found in 3 cats, while the others were only found in a single cat each. In the previous

study (Brown et al., 2009a), infection with ITSa was significantly associated with a fatal clinical

outcome but unfortunately, the clinical outcome of the cats from Brown et al., (2009b) could not

be determined because many of the cats were euthanized and submitted as clinical cases. The

genotype ITSA from the previous study was only identified in 3 samples from different years,

which supports the previous findings that this genotype may be rare in eastern states (Brown et

al. 2009b).

Further studies are required from more spatially diverse domestic cat populations to

determine if specific genotypes are circulating in domestic cats. Currently the data support that

certain genotype are associated with specific geographic areas (Brown et al., 2009b).

Furthermore, one particular genotype has been associated with pathogenicity for domestic cats

(Brown et al., 2009a). Whether or not the strains of C. felis identified in these studies reflect the

strains of C. felis which circulate in wild felids or the tick vectors is unknown.

28

Cytauxzoon spp. from felid species outside of the United States

Germany

The only report of Cytauxzoon in Germany was a fatal infection in 1984 in a female

Bengal tiger (Panthera tigris) which was captive-born in a German zoo (Jakob and Wesemeier,

1996). The tiger was 18 weeks old when it became anorexic, lethargic, and dyspnoeic and

rapidly died from acute pneumonia (Jakob and Wesemeier, 1996). Gross findings revealed

heavily congested lungs with numerous petechial hemorrhages. All of the examined lymph nodes

and the spleen were enlarged and hemorrhagic. Microscopic evaluation revealed mononuclear

phagocytes occluding the blood vessels of all examined organs. Macroschizonts were clearly

visible. Piroplasms were observed in the erythrocytes. Cytauxzoonosis is suspected based on the

congestion and edema of the lungs, splenomegaly and hemorrhagic lymph nodes, and

observation of intramononuclear cell schizonts. Unfortunately genetic characterization of the

samples was not conducted. No ticks were observed on the tiger and a brother to the tiger was

born and raised to maturity in the same enclosure. Importantly, 14 months before the death of the

young tiger, three bobcats had been imported directly from the United States to the zoo.

Parasitemia has been shown to last over four months in bobcats, and many reports suggest that

the animals may remain subclinically infected for years (Keir et al., 1982b). At the time, no

examinations were conducted on the bobcats or their blood, so the origin of the infection is

unknown (Jakob and Wesemeier, 1996). As many C. felis researchers have indicated (Millan et

al., 2007; Reichard et al., 2005), transport of wild felids without a full clinical evaluation and

quarantine period can have devastating consequences.

29

Spain

The first report of a Cytauxzoon sp. in Europe was in a domestic cat from Spain (Criado-

Fornelio et al., 2004). Only a single cat of 100 (1%) was PCR positive for a Cytauxzoon sp. that

was, based on 18S rRNA gene sequence analysis, 98% similar to C. manul and 95% with a

Cytauxzoon sp. (stabilate 153) that was originally reported to be from a domestic cat from South

Africa (Criado-Fiorello et al., 2004), however stabiliate 153 is believed to be from a cat from

The Netherlands that was subcutaneously inoculated with a domestic cat spleen stabiliate from

Missouri, USA (Uilenberg et al., 1987). Phylogenetic analysis of the Cytauxzoon sp. from Spain

(see phylogenetic tree, Figure 1.1) suggests that it is a distinct species from C. felis.

The first report of a Cytauxzoon sp. in a wild European felid was in the Iberian lynx

(Lynx pardinus) which is one of the world’s most endangered felid and is only found in two

isolated populations in southern Spain (Nowell and Jackson, 1996; Johnson et al., 2004; Peña et

al., 2003; Millan et al., 2007). The Doñana National Park population has approximately 40

individuals while the Sierra Morena population contains between 90 and 120 individuals

(Guzman et al., 2003). The first report was from a free-ranging injured lynx that was taken for a

health check from the Sierra Morena population and examination of a blood smear revealed a 4%

parasitemia. Sequence analysis of 1,675 bp of the 18S rRNA gene indicated that the Cytauxzoon

sp. from the lynx only differed from the domestic cat Spanish sample by 11 bases. Interestingly,

the lynx Cytauxzoon sp. was 99.6% identical to C. manul (differed by only 8 bases) (Luaces et

al., 2005). A retrospective analysis of blood and tissue samples collected from 50 lynx from

1993-2003 was uniformly negative for Cytauxzoon infections (Luaces et al., 2005). Since no

clinical signs were observed in the infected male lynx from this study, the infection was assumed

to be subclinical. Nevertheless, this Cytauxzoon sp. is a concern because the Iberian lynx is

30

endangered and typically nonpathogenic organisms can become pathogenic due to inbreeding

and during coinfections with pathogens that cause immunodeficiency, the Cytauxzoon sp. is a

serious consideration (Johnson et al., 2004; Peña et al., 2003).

A study on the prevalence of the Cytauxzoon sp. in wild-caught lynx from Sierra Morena

found that 3 of 9 (33%) lynx were infected (Millan et al., 2007). All three were juveniles. Eleven

lynx from Doñana were all negative. Sequence analysis of the 18S rRNA gene of these positives

revealed minimal variation (differed from each other by 0.6%, but differed from the previous

lynx sample by 1%). According to this analysis, the sequences were 99.4% identical to C. felis

from the United States. Currently no vector has been identified and the lack of infection in the

Doñana population may be due to lack of vector either because of environmental restrictions or

host-specific extinction.

France

Cytauxzoon sp. infections of domestic cats in France were first reported in 2008 (Criado-

Fornelio et al., 2008). A single cat of 116 cats (0.8%) surveyed were PCR positive for a

Cytauxzoon sp. Genetic characterization of the 18S rRNA region revealed that the piroplasm was

99% similar to the Cytauxzoon sp. described in Spanish felids (Criado-Fornelio et al., 2004;

Criado-Fornelio et al., 2007b; Millan et al., 2007). The low prevalence in domestic cats in France

was similar to the low prevalence in domestic cats in Spain (Criado-Fornelio et al., 2004; Criado-

Fornelio et al., 2007b).

31

Mongolia

In October and December of 2000, four Pallas’s cats (Otocolobus manul) were

transported from Mongolia to Oklahoma. During route quarantine examination, small

intraerythrocytic piroplasms were observed on stained blood smears and parasitemias <1%.

Morphologic analysis and host (felid) suggested that the piroplasms were C. felis. Amplification

and sequencing of the 1,485 bp of the rRNA gene indicated that it was related to C. felis, but

likely represented a different species of Cytauxzoon (only 3.6% different from C. felis compared

to the differences from Babesia leo (10.13%) and B. felis (9.49%)). This was the first report of

erythroparasitemia in Pallas’s cats and probably the first report of a piroplasm in free-ranging

felids in Mongolia (Ketz-Riley et al., 2003).

The initial study of the Cytauxzoon sp. in Mongolian Pallas’s cats did not formally

propose a new species for the organism. In 2005, a second study was conducted to further

investigate the phylogenetic relationship between this Cytauxzoon sp. and C. felis (Reichard et

al., 2005). In the first study, only blood from one of the four imported Pallas’s cats was available

for analysis, but in the second study, samples from two of the remaining cats were analyzed.

After PCR amplification and sequencing of the entire18S rRNA gene, the authors concluded that

the mean corrected percent sequence divergence between the Pallas’s cat Cytauxzoon sp. and C.

felis was 1.49%. In addition, the Pallas’s cat Cytauxzoon sp. was found to only be 0.389%

divergent from a Spanish sample of a Cytauxzoon-like sp.. The species name Cytauxzoon manul,

based on the vertebrate host from which the piroplasm was first detected, was proposed

(Reichard et al., 2005). The vector for C. manul is currently unknown, but is presumably an

Ixodid tick.

32

Recently, a random, blind study was conducted to determine both the infectivity of C.

manul for domestic cats as well as the ability of C. manul to provide cross-protection for

infection with C. felis (Joyner et al., 2007). Three cats were immunosuppressed with

dexamethasone and three others were inoculated with sterile water, after which, the cats were

intravenously inoculated with 2 ml of heparinized blood from the i-positive Pallas’s cat blood

(parasitemia >1x106). None of the cats displayed clinical signs of disease, but both groups

became parasitemic within two weeks of inoculation and at least two cats were still parasitemic

37 days post-inoculation. Weekly splenic aspirates were performed and no schizogenous forms

of C. manul were found. All of the domestic cats were then challenged with C. felis-infected

splenic homogenate and all developed clinical cytauxzoonosis within 5 days post-inoculation.

The findings from this study suggest that the intraerythrocytic stage of C. manul is infective for

domestic cats, but that the tissue stage of the infection does not occur. Importantly, cat inoculated

with C. manul did not develop any cross-protection against subsequent C. felis infection.

The lack of schizongony has been reported in other studies involving C. felis and bobcats and

domestic cats. When domestic cats and bobcats are incoculated with intraerythrocytic states of C.

felis, disease is limited, but when bobcats or domestic cats are inoculated with schizogenous

stages, severe disease can result (Glenn et al., 1983; Kier et al., 1982b). It is unknown if clinical

disease would have developed in domestic cats if they had been inoculated with schizogenous

stages of C. manul (Joyner et al., 2007).

Brazil

A piroplasm, most likely a Cytauxzoon sp., was first reported in Brazil in 2004 a colony

of urban stray domestic cats living in the Rio de Janeiro zoological garden (Mendes-de-Almeida

33

et al., 2004). The general goal of the project was to understand the pathogens and other health

problems associated with feral domestic cats. Over a two month period, 47 cats were captured

for the study and full genetic and health evaluations were conducted. Besides finding other

pathogens such as mycoplasmas (Haemobartonella felis, 38%), the authors discovered a small

intraerythrocytic piroplasm in 47% of the cats. Based on morphology, the authors were unable to

identify the hemoparasite as either a Babesia sp. or a Cytauxzoon sp. No ticks were found on any

of the animals. To date, no phylogenetic analysis has been conducted on the specimens to

determine their identity, but this was the first report of a felid piroplasm in South America

(Mendes-de-Almeida et al., 2004).

Fatal cytauxzoonosis was reported in a captive reared lioness (Panthera leo) and her 6-

month-old cub in the Volta Redonda Municipal Zoo, Rio de Janeiro State, Brazil (Peixoto et al.,

2007). In 1998, the 6-month old cub died after a 24-hour depression. Gross examination revealed

hemothorax, endocardial and pulmonary edema, and petechial hemorrhages. Forty-five days later

the mother of the cub also died. Clinical signs were observed five days before its death. The

clinical signs and lesions in the lioness were suggestive of cytauxzoonosis including pulmonary

edema, depression, anemia, thrombocytopenia and petechial hemorrhages. Several biochemical

abnormalities including pyuria, hematuria, proteinuria, as well as pulmonary edema and

petechial hemorrhages were observed in the lioness. Microscopic evaluation of tissues from both

felids revealed mononuclear phagocytes containing schizogenous forms consistent with a

Cytauxzoon sp. Electron microscopy revealed schizonts consistent with Cytauxzoon which were

delimited by a thin double membrane inside the cytoplasm of macrophages. Amblyomma

cajennense ticks were collected from the lioness. The authors believed that this infection

represented a South American strain of C. felis because the organisms observed were

34

ultrastructrually identical to C. felis and sequence analysis of the 18S rRNA region from both

these samples found a 99% similarity with C. felis from a domestic cat from the United States

(Scofield, 2006). This study represents the first report of a Cytauxzoon sp. infection in P. leo.

In 2006, blood samples were collected from 72 Brazilian wild-captive felids from five

zoos and screened for infection with Cytauxzoon sp. (Andre et al., 2009). There were seven felid

species included in the study including: 9 pumas (P. concolor); 29 ocelots (Leopardus pardalis);

6 jaguarondi (Puma yagouaroundi); 2 margays (Leopardus wiedii); 14 little spotted cats

(Leopardus tigrinus); 3 pampas cats (Oncifelis colocolo); and 9 jaguars (Panthera onca). PCR

testing revealed that 13% of the animals were infected, including six ocelots, two pumas, and one

jaguar from two different zoos. The Cytauxzoon spp. detected in these felids was 99% similar to

the previous sample from the Brazilian lioness and 98% similar to C. felis from domestic cats.

No clinical signs or symptoms were associated with the infection in these felids and no ticks

were found on the felids. The authors concluded that wild felids in Brazil, similar to wild felids

in the United States, may be reservoirs for Cytauxzoon species. And, like the occasional death in

bobcats in the United States (Nietfeld and Pollack, 2002), native wild felids in Brazil may

occasionally die from infection (Peixoto et al., 2007).

Africa

At least two papers have reported Cytauxzoon-like organisms from wild felids in Africa

and an additional two papers from Africa have reported piroplasms indistinguishable from

Cytauxzoon or Babesia. In 1929, Davis described a piroplasm of the Sudanese wild cat (Felis

ocreata, probably Felis silvestris), but it unclear if schizogenous stages were observed and no

further studies have been conducted. Piroplasms have also been described from the West African

35

civet cat (Viverra civetta), but no further analyses have been conducted (Wenyon and Hamerton,

1930). Cytauxazoon-like organisms in the erythrocytes of two cheetahs from Africa have been

reported (Zindle et al., 1981). In 1982, a Cytauxzoon-like disease was reported in a domestic cat

from Zimbabwe (Foggin and Roberts, 1982). Based on necropsy, the cat had schizogenous stages

in reticuloendothelial cells in the spleen, lymph nodes, lungs and liver. The researchers stated

that this disease had been observed in other domestic felids from the area. In a separate study,

piroplasms were observed in erythrocytes of genets (Genetta genetta). A domestic cat was

inoculated with blood from an infected genet and developed parasitemia. Then the cat was

treated with imidocarb and survived and no necropsy was performed (Foggin and Roberts,

1982). Although no further studies have been conducted on wild or domestic felids in Zimbabwe,

this evidence strongly suggests that Cytauxzoon is circulating in African felids.

LITERATURE CITED

Allan, S.A., In: Samuel, W.M., Pybus, M.J., Kocan, A.A. (Eds.), Parasitic Diseases of Wild

Mammals. Iowa State University Press, Ames, Iowa, 72-94.

Allsopp, M.T., Lewis, B.D., Penzhorn, B.L., 2007, Molecular evidence for transplacental

transmission of Theileria equi from carrier mares to their apparently healthy foals. Vet

Parasitol 148, 130-136.

Andre, M.R., Adania, C.H., Machado, R.Z., Allegretti, S.M., Felippe, P.A., Silva, K.F., Nakaghi,

A.C., Dagnone, A.S., 2009, Molecular detection of Cytauxzoon spp. in asymptomatic

Brazilian wild captive felids. J Wildl Dis 45, 234-237.

36

Baek, B.K., Soo, K.B., Kim, J.H., Hur, J., Lee, B.O., Jung, J.M., Onuma, M., Oluoch, A.O.,

Kim, C.H., Kakoma, I., 2003, Verification by polymerase chain reaction of vertical

transmission of Theileria sergenti in cows. Can J Vet Res 67, 278-282.

Baneth, G., Kenny, M.J., Tasker, S., Anug, Y., Shkap, V., Levy, A., Shaw, S.E., 2004, Infection

with a proposed new subspecies of Babesia canis, Babesia canis subsp. presentii, in

domestic cats. J Clin Microbiol 42, 99-105.

Barta, J.R., Coles, B.A., Schito, M.L., Fernando, M.A., Martin, A., Danforth, H.D., 1998,

Analysis of infraspecific variation among five strains of Eimeria maxima from North

America. Int J Parasitol 28, 485-492.

Bendele, R.A., Schwartz, W.L., Jones, L.P.,1976, Cytauxzoon-like disease in Texas cats.

Southwest Vet 29, 244-246.

Bigalke, R.D., 1989, The current status of research on diseases of wildlife in South Africa and

south west Africa/Namibia. J S Afr Vet Assoc 60, 7-10.

Birkenheuer, A.J., Cohn, L.A., Levy, M.G., Breitschwerdt, E.B., Marr, H.S., 2008a, Atovaquone

and azithromycin for the treatment of Cytauxzoon felis. J Vet Intern Med 22, 703-704.

Birkenheuer, A.J., Le, J.A., Valenzisi, A.M., Tucker, M.D., Levy, M.G., Breitschwerdt, E.B.,

2006a, Cytauxzoon felis infection in cats in the mid-Atlantic states: 34 cases (1998-2004).

J Am Vet Med Assoc 228, 568-571.

Birkenheuer, A.J., Marr, H., Alleman, A.R., Levy, M.G., Breitschwerdt, E.B., 2006b,

Development and evaluation of a PCR assay for the detection of Cytauxzoon felis DNA

in feline blood samples. Vet Parasitol 137, 144-149.

37

Birkenheuer, A.J., Marr, H.S., Warren, C., Acton, A.E., Mucker, E.M., Humphreys, J.G., Tucker,

M.D., 2008b, Cytauxzoon felis infections are present in bobcats (Lynx rufus) in a region

where cytauxzoonosis is not recognized in domestic cats. Vet Parasitol 153, 126-130.

Blouin, E.F., Kocan, A.A., Glenn, B.L., Kocan, K.M., Hair, J.A., 1984, Transmission of

Cytauxzoon felis Kier, 1979 from bobcats, Felis rufus (Schreber), to domestic cats by

Dermacentor variabilis (Say). J Wildl Dis 20, 241-242.

Blouin, E.F., Kocan, A.A., Kocan, K.M., Hair, J., 1987, Evidence of a Limited Schizogonous

Cycle for Cytauxzoon felis in Bobcats Following Exposure to Infected Ticks. J Wildlife

Dis 23, 499-501.

Bondy, P.J., Cohn, L.A., Kerl, M.E., 2005a, Feline cytauxzoonosis. Comp Cont Educ Pract 27,

69-75.

Bondy, P.J., Jr., Cohn, L.A., Tyler, J.W., Marsh, A.E., 2005b, Polymerase chain reaction

detection of Cytauxzoon felis from field-collected ticks and sequence analysis of the

small subunit and internal transcribed spacer 1 region of the ribosomal RNA gene. J

Parasitol 91, 458-461.

Brocklesby, D.W., 1962, Cytauxzoon taurotragi (Martin and Brocklesby, 1960) a piroplasm of

the eland (Taurotragus oryx pattersonianus (Lydekker, 1906)). Research in Vet Sci 3,

334-344.

Brocklesby, D.W., 1979, Recent observations on tick-borne protozoa. In: Wilde J.K.H. (ed)

Tick-Borne Diseases and their Vectors. Edinburgh University Press, 262-286.

Brown, H.M., Berghaus, R.D., Latimer, K.S., Britt, J.O., Rakich, P.M., Peterson, D.S., 2009,

Genetic variability of Cytauxzoon felis from 88 infected domestic cats in Arkansas and

Georgia. J Vet Diagn Invest 21, 59-63.

38

Brown, H.M., Modaresi, S.M., Cook, J.L., Latimer, K.S., Peterson, D.S., 2009, Genetic

variability of archived Cytauxzoon felis histologic specimens from domestic cats in

Georgia, 1995-2007. J Vet Diagn Invest 21, 493-498.

Brown, H.M., Latimer, K.S., Erikson, L.E., Cashwell, M.E., Britt, J.O., Peterson, D.S., 2008,

Detection of persistent Cytauxzoon felis infection by polymerase chain reaction in three

asymptomatic domestic cats. J Vet Diagn Invest 20, 485-488.

Butt, M.T., Bowman, D., Barr, M.C., Roelke, M.E., 1991, Iatrogenic transmission of Cytauxzoon

felis from a Florida panther (Felix concolor coryi) to a domestic cat. J Wildl Dis 27, 342-

347.

Carmichael, I.H., Hobday, E., 1975, Blood parasites of some wild bovidae in Botswana.

Onderstepoort J Vet Res 42, 55-62.

Cowell, R.L., Fox, J.C., Panciera, R.J., Tyler, R.D., 1988, Detection of Anticytauxzoon

Antibodies in Cats Infected with a Cytauxzoon Organism from Bobcats. Vet Parasitol 28,

43-52.

Criado-Fornelio, A., 2007, A review of nucleic-acid-based diagnostic tests for Babesia and

Theileria, with emphasis on bovine piroplasms. Parassitologia 49 Suppl 1, 39-44.

Criado-Fornelio, A., Buling, A., Pingret, J.L., Etievant, M., Boucraut-Baralon, C., Alongi, A.,

Agnone, A., Torina, A., 2009, Hemoprotozoa of domestic animals in France: Prevalence

and molecular characterization. Vet Parasitol 159, 73-76.

Criado-Fornelio, A., Gonzalez-del-Rio, M.A., Buling-Sarana, A., Barba-Carretero, J.C., 2004,

The "expanding universe" of piroplasms. Vet Parasitol 119, 337-345.

39

Criado-Fornelio, A., Martinez-Marcos, A., Buling-Sarana, A., Barba-Carretero, J.C., 2003a,

Molecular studies on Babesia, Theileria and Hepatozoon in southern Europe. Part I.

Epizootiological aspects. Vet Parasitol 113, 189-201.

Criado-Fornelio, A., Martinez-Marcos, A., Buling-Sarana, A., Barba-Carretero, J.C., 2003b,

Molecular studies on Babesia, Theileria and Hepatozoon in southern Europe. Part II.

Phylogenetic analysis and evolutionary history. Vet Parasitol 114, 173-194.

Criado-Fornelio, A., Martinez-Marcos, A., Buling-Sarana, A., Barba-Carretero, J.C., 2003c,

Presence of Mycoplasma haemofelis, Mycoplasma haemominutum and piroplasmids in

cats from southern Europe: a molecular study. Vet Microbiol 93, 307-317.

Davis, 1929, On a piroplasm of the Sudanese wild cat (Felis ocreata), Trans Roy Soc Trop Med

Hyg 22, 523-534.

Edwards, A.C., Reichard, M.V., Meinkoth, J.H., Snider, T.A., Meinkoth, K.R., Heinz, R.E.,

Little, S.E., 2010, Confirmation of Amblyomma americanum (Acari: Ixodidae) as a vector

for Cytauxzoon felis (Piroplasmorida: Theileriidae) to domestic cats. J Med Ent.

Ferris, D.H., 1979, A progress report on the status of a new disease of American cats:

cytauxzoonosis. Comp Immunol Microbiol Infect Dis 1, 269-276.

Foggin, C.M., Roberts, H.M., 1982, A cytauxzoonosis-like disease in a cat in Zimbabwe,

Zimbabwe Vet J 13, 28-29.

Garner, M.M., Lung, N.P., Citino, S., Greiner, E.C., Harvey, J.W., Homer, B.L., 1996, Fatal

cytauxzoonosis in a captive-reared white tiger (Panthera tigris). Vet Pathol 33, 82-86.

Glenn, B.L., Kocan, A.A., Blouin, E.F., 1983, Cytauxzoonosis in bobcats. J Am Vet Med Assoc

183, 1155-1158.

40

Glenn, B.L., Rolley, R.E., Kocan, A.A., 1982, Cytauxzoon-like piroplasms in erythrocytes of

wild-trapped bobcats in Oklahoma. J Am Vet Med Assoc 181, 1251-1253.

Glenn, B.L., Stair, E.L., 1984, Cytauxzoonosis in domestic cats: report of two cases in

Oklahoma, with a review and discussion of the disease. J Am Vet Med Assoc 184, 822-

825.

Greene, C.E., Latimer, K., Hopper, E., Shoeffler, G., Lower, K., Cullens, F., 1999,

Administration of diminazene aceturate or imidocarb dipropionate for treatment of

cytauxzoonosis in cats. J Am Vet Med Assoc 215, 497-500, 482.

Guzman, J.N., Garcia, F.J., Garrote, G., Perez de Ayala, R., Iglesias, M.C., 2003, Resultados del

censo diagnostico de las poblaciones de lince iberico (Lynx pardinus) en Espana (2002).

In proceedings: VI Jornadas de la Sociedad Espanola para la Conservacion y el Estudio

de los Mamiferos- an Iberian Conference, Secretaria de la SECEM (ed.). SECEM, Apdo.

15450, 29080, Malaga, Spain, 89.

Haber, M.D., Tucker, M.D., Marr, H.S., Levy, J.K., Burgess, J., Lappin, M.R., Birkenheuer, A.J.,

2007, The detection of Cytauxzoon felis in apparently healthy free-roaming cats in the

USA. Vet Parasitol 146, 316-320.

Harvey, J.W., Dunbar, M.R., Norton, T.M., Yabsley, M.J., 2007, Laboratory findings in acute

Cytauxzoon felis infection in cougars (Puma concolor couguar) in Florida. J Zoo Wildlife

Med 38, 285-291.

Hauck, W.N., Snider, T.G., Lawrence, J.E., 1982, Cytauxzoonosis in a native Louisiana cat, J

Am Vet Med Asso 180, 1472-1474.

41

Hazen-Karr, C.G., Kocan, A.A., Kocan, K.M., Hair, J.A., 1987, The ultrastructure of sporogony

in Theileria cervi (Bettencourt et al., 1907) in salivary glands of female Amblyomma

americanum (L.) ticks. J Parasitol 73, 1182-1188.

Hillis, D.M., Dixon, M.T., 1991, Ribosomal DNA: molecular evolution and phylogenetic

inference. Q Rev Biol 66, 411-453.

Hoover, J.P., Walker, D.B., Hedges, J.D., 1994, Cytauxzoonosis in cats: eight cases (1985-1992).

J Am Vet Med Assoc 205, 455-460.

Jackson, C.B., Fisher, T., 2006, Fatal cytauxzoonosis in a Kentucky cat (Felis domesticus). Vet

Parasitol 139, 192-195.

Jacobson, L.S., Schoeman, T., Lobetti, R.G., 2000, A survey of feline babesiosis in South Africa.

J S Afr Vet Assoc 71, 222-228.

Jakob, W., Wesemeier, H.H., 1996, A fatal infection in a Bengal tiger resembling

cytauxzoonosis in domestic cats. J Comp Pathol 114, 439-444.

Jardine, J.E., 1992, The pathology of cytauxzoonosis in a tsessebe (Damaliscus lunatus). J S Afr

Vet Assoc 63, 49-51.

Joyner, P.H., Reichard, M.V., Meinkoth, J.H., Milne, V.E., Confer, A.W., Kocan, A.A., Hoover,

J.P., 2007, Experimental infection of domestic cats (Felis domesticus) with Cytauxzoon

manul from Pallas' cats (Otocolobus manul). Vet Parasitol 146, 302-306.

Keirans, J.E., Lacombe, E.H., 1998, First records of Amblyomma americanum, Ixodes (Ixodes)

dentatus, and Ixodes (Ceratixodes) uriae (Acari: Ixodidae) from Maine. J Parasitol 84,

629-631.

42

Ketz-Riley, C.J., Reichard, M.V., Van Den Bussche, R.A., Hoover, J.P., Meinkoth, J., Kocan,

A.A., 2003, An intraerythrocytic small piroplasm in wild-caught Pallas's cats

(Otocolobus manul) from Mongolia. J Wildlife Dis 39, 424-430.

Kier, A.B., Wagner, J.E., Kinden, D.A., 1987, The pathology of experimental cytauxzoonosis. J

Comp Pathol 97, 415-432.

Kier, A.B., Wagner, J.E., Morehouse, L.G., 1982a, Experimental Transmission of Cytauxzoon

felis from Bobcats (Lynx rufus) to Domestic Cats (Felis domesticus). Am J Vet Res 43,

97-101.

Kier, A.B., Wightman, S.R., Wagner, J.E., 1982b, Interspecies transmission of Cytauxzoon felis.

Am J Vet Res 43, 102-105.

Kocan, A.A., Blouin, E.F., Glenn, B.L., 1985, Hematologic and serum chemical values for free-

ranging bobcats, Felis rufus (Schreber), with reference to animals with natural infections

of Cytauxzoon felis Kier, 1979. J Wildlife Dis 21, 190-192.

Kocan, A.A., Kjemtrup, A., Meinkoth, J., Whitworth, L.C., Murphy, G.L., Decker, L., Lorenz,

M., 2001, A genotypically unique Babesia gibsoni-like parasite recovered from a dog in

Oklahoma. J Parasitol 87, 437-438.

Kocan, A.A., Kocan, K.M., 1991, Tick-transmitted diseases of wildlife in North America. Bull

Soc Vector Eco 16, 94-108.

Kocan, A.A., Kocan, K.M., Blouin, E.F., Mukolwe, S.W., 1992, A Redescription of schizogony

of Cytauxzoon felis in the domestic cat. Ann Ny Acad Sci 653, 161-167.

Kocan, K.M., Wickwire, K.B., Ewing, S.A., Hair, J.A., Barron, S.J., 1988, Preliminary studies of

the development of Anaplasma marginale in salivary glands of adult, feeding

Dermacentor andersoni ticks. Am J Vet Res 49, 1010-1013.

43

Kocan, A.A., Waldrup, K.A., 2001, In: Samuel, W.M., Pybus, M.J., Kocan, A.A. (Eds.),

Parasitic Diseases of Wild Mammals. Iowa State University Press, Ames, Iowa, 524-536.

Koch, H.G., Dunn, J.C., 1980a, Ticks collected from small and medium-sized wildlife hosts in

LeFlore County, Oklahoma. Southwestern Ent 5, 214-221.

Levine, N.D., Corliss, J.O., Cox, C.E., Deroux, G., Grain, J., Honigberg, B.M., Leedale, G.F.,

Loeblich, A.R., Lom, J., Lynn, D., Merinfeld, E.G., Page, F.C., Poljansky, G., Sprague,

V., Vavra, J., Wallace, F.G., 1980, A newly revised classification of the protozoa. J

Protozoology 27, 37-58.

Lopez-Rebollar, L.M., Penzhorn, B.L., de Waal, D.T., Lewis, B.D., 1999, A possible new

piroplasm in lions from the Republic of South Africa. J Wildlife Dis 35, 82-85.

Luaces, I., Aguirre, E., Garcia-Montijano, M., Velarde, J., Tesouro, M.A., Sanchez, C., Galka,

M., Fernandez, P., Sainz, A., 2005, First report of an intraerythrocytic small piroplasm in

wild Iberian lynx (Lynx pardinus). J Wildlife Dis 41, 810-815.

McCully, R.M., Keep, M.E., Basson, P.A., 1970, Cytauxzoonosis in a giraffe (Giraffa

camelopardalis (Linnaeus, 1758)) in Zululand. Onderstepoort J Vet Res 37, 7-9.

Meier, H.T., Moore, L.E., 2000, Feline cytauxzoonosis: A case report and literature review. J

Am Anim Hosp Assoc 36, 493-496.

Meinkoth, J., Kocan, A.A., Whitworth, L., Murphy, G., Fox, J.C., Woods, J.P., 2000, Cats

surviving natural infection with Cytauxzoon felis: 18 cases (1997-1998). J Vet Intern Med

14, 521-525.

Meinkoth, J.H., Kocan, A.A., 2005, Feline cytauxzoonosis. Vet Clin N Am-Small 35, 89.

Mendes-de-Almeida, F., Faria, M.C., Branco, A.S., Serrao, M.L., Souza, A.M., Almosny, N.,

Charme, M., Labarthe, N., 2004, Sanitary conditions of a colony of urban feral cats (Felis

44

catus Linnaeus, 1758) in a zoological garden of Rio de Janeiro, Brazil. Rev Inst Med

Trop Sao Paulo 46, 269-274.

Millan, J., Naranjo, V., Rodriguez, A., de la Lastra, J.M., Mangold, A.J., de la Fuente, J., 2007,

Prevalence of infection and 18S rRNA gene sequences of Cytauxzoon species in Iberian

lynx (Lynx pardinus) in Spain. Parasitology 134, 995-1001.

Motzel, S.L., Wagner, J.E., 1990, Treatment of experimentally induced cytauxzoonosis in cats

with parvaquone and buparvaquone. Vet Parasitol 35, 131-138.

Neitz, W.O., Thomas, A.D., 1948, Cytauxzoon sylvicaprae gen. nov., spec. nov., a protozoon

responsible for a hitherto undescribed disease in the duiker, Sylvicapra grimmia (Linne).

Onderstepoort J Vet Sci Anim Ind 23, 63-76.

Neitz, W.O., 1957, Theileriosis, gonderioses and cytauxzoonoses. A review. Onderstepoort J Vet

Res 27, 275-430.

Neitz, W.O., 1964, Cytauxzoonosis in African antelopes, Bulletin de l'Office International des

Epizooties 62, 1009-1015.

Nietfeld, J.C., Pollock, C., 2002, Fatal cytauxzoonosis in a free-ranging bobcat (Lynx rufus). J

Wildlife Dis 38, 607-610.

Nijhof, A.M., Pillay, V., Steyl, J., Prozesky, L., Stoltsz, W.H., Lawrence, J.A., Penzhorn, B.L.,

Jongejan, F., 2005, Molecular characterization of Theileria species associated with

mortality in four species of African antelopes. J Clin Microbiol 43, 5907-5911.

Nowell, K., Jackson, P., 1996, Wild cats: status survey and conservation action plan. IUCN,

Gland, Switzerland.

45

Olivier, C., van de Pas, S., Lepp, P.W., Yoder, K., Relman, D.A., 2001, Sequence variability in

the first internal transcribed spacer region within and among Cyclospora species is

consistent with polyparasitism. Int J Parasitol 31, 1475-1487.

Peixoto, P.V., Soares, C.O., Scofield, A., Santiago, C.D., Franca, T.N., Barros, S.S., 2007, Fatal

cytauxzoonosis in captive-reared lions in Brazil. Vet Parasitol 145, 383-387.

Pena, L., Garcia, P., Fernandez, P., Galka, M., Benito, A., Sanchez, C., 2003, Histological

evidences of immunodeficiency in Iberian lynx (Lynx pardinus). In Proceedings: 21st

Annual Conference of the European Society of Veterinary Pathologists, Dublin.

Penzhorn, B.L., 2006, Babesiosis of wild carnivores and ungulates. Vet Parasitol 138, 11-21.

Penzhorn, B.L., Schoeman, T., Jacobson, L.S., 2004, Feline babesiosis in South Africa: a review.

Ann N Y Acad Sci 1026, 183-186.

Phipps, L.P., Otter, A., 2004, Transplacental transmission of Theileria equi in two foals born and

reared in the United Kingdom. Vet Rec 154, 406-408.

Reichard, M.V., Baum, K.A., Cadenhead, S.C., Snider, T.A., 2008a, Temporal occurrence and

environmental risk factors associated with cytauxzoonosis in domestic cats. Vet Parasitol

152, 314-320.

Reichard, M.V., Meinkoth, J.H., Edwards, A.C., Snider, T.A., Kocan, K.M., Blouin, E.F., Little,

S.E., 2008b, Transmission of Cytauxzoon felis to a domestic cat by Amblyomma

americanum. Vet Parasitol.

Reichard, M.V., Meinkoth, J.H., Edwards, A.C., Snider, T.A., Kocan, K.M., Blouin, E.F., Little,

S.E., 2009, Transmission of Cytauxzoon felis to a domestic cat by Amblyomma

americanum. Vet Parasitol 161, 110-115.

46

Reichard, M.V., Van Den Bussche, R.A., Meinkoth, J.H., Hoover, J.P., Kocan, A.A., 2005, A

new species of Cytauxzoon from Pallas' cats caught in Mongolia and comments on the

systematics and taxonomy of piroplasmids. J Parasitol 91, 420-426.

Rotstein, D.S., Taylor, S.K., Harvey, J.W., Bean, J., 1999, Hematologic effects of

cytauxzoonosis in Florida panthers and Texas cougars in Florida. J Wildlife Dis 35, 613-

617.

Schoeman, T., Lobetti, R.G., Jacobson, L.S., 2001, Feline babesiosis: signalment, clinical

pathology, and co-infection. J S Afr Vet Assoc 72, 4-11.

Scofield, A., 2006, Analises genotipica e filogenetica do gene 18S RNAr de isolados brasileiros

de Cytauxzoon felis (Apicomplexa: Theileriidae), Thesis, Universidad Federal Rural do

Rio de Janeiro, Brazil.

Semtner, P.J., Hair, J.A., 1973a, The ecology and behavior of the lone star tick (Acarina:

Ixodidae). IV. The daily and seasonal activity patterns of adults in different habitat types.

J Med Entomol 10, 337-344.

Semtner, P.J., Hair, J.A., 1973b, The ecology and behavior of the lone star tick (Acarina:

Ixodidae). V. Abundance and seasonal distribution in different habitat types. J Med

Entomol 10, 618-628.

Semtner, P.J., Sauer, J.R., Hair, J.A., 1973, The ecology and behavior of the lone star tick

(Acarina: Ixodidae). 3. The effect of season on molting time and post-molt behavior of

engorged nymphs and adults. J Med Entomol 10, 202-205.

Shindel, N., Dardiri, A.H., Ferris, D.H., 1978, Indirect Fluorescent-Antibody Test for Detection

of Cytauxzoon-Like Organisms in Experimentally Infected Cats. Can J Comp Med 42,

460-465.

47

Simpson, C.F., Harvey, J.W., Carlisle, J.W., 1985, Ultrastructure of the intraerythrocytic stage of

Cytauxzoon felis. Am J Vet Res 46, 1178-1180.

Snider, T.A., Confer, A.W., Payton, M.E., 2010, Pulmonary histopathology of Cytauxzoon felis

infections in the cat. Vet Pathol. (In press).

Sonenshine, D.E., Levy, G.F., 1971, The ecology of the lone star tick, Amblyomma americanum

(L.), in two contrasting habitats in Virginia (Acarina: Ixodidae). J Med Entomol 8, 623-

635.

Susta, L., Torres-Velez, F., Zhang, J., Brown, C., 2009, An in situ hybridization and

immunohistochemical study of cytauxzoonosis in domestic cats. Vet Pathol 46, 1197-

1204.

Uilenberg, G., Franssen, F.F., Perie, N.M., 1987, Relationships between Cytauxzoon felis and

African piroplasmids. Vet Parasitol 26, 21-28.

Wagner, J.E., 1976, A fatal cytauxzoonosis-like disease in cats. J Am Vet Med Assoc 168, 585-

588.

Wagner, J.E., Morehouse, L.G., 1976, Animal susceptibility studies with a Cytauxzoon agent of

feline origin. J Am Vet Med Assoc 169, 1135-1135.

Walker, D.B., Cowell, R.L., 1995, Survival of a domestic cat with naturally acquired

cytauxzoonosis. J Am Vet Med Assoc 206, 1363-1365.

Weisman, J.L., Woldemeskel, M., Smith, K.D., Merrill, A., Miller, D., 2007, Blood smear from a

pregnant cat that died shortly after partial abortion. Vet Clin Path 36, 209-211.

Wenyon, C.M., Hamerton, A.E., 1930, Piroplasms of the West African civet cat (Viverra civetta)

and the Bay lynx (Felis rufa) of North America. Roy Soc Trop Med and Hyg, 24, 7-8.

48

Wightman, S.R., Kier, A.B., Wagner, J.E., 1977, Feline cytauxzoonosis: clinical features of a

newly described blood parasite disease. Feline Pract, 24-26.

Wilson, D.E., Bartsch, R.C., Bigalke, R.D., Thomas, S.E, 1974, Observations on mortality rates

and disease in roan and sable antelope in nature reserves in the Transvall. J S African

Wildl Manag Asso 4, 203-206.

Wilson, D.E., Hirst, S.M., 1977, Ecology and factors limiting roan and sable antelope

populations in South Africa, Wildl Monographs 54.

Yabsley, M.J., Murphy, S.M., Cunningham, M.W., 2006, Molecular detection and

characterization of Cytauxzoon felis and a Babesia species in cougars from Florida. J

Wildlife Dis 42, 366-374.

Zahler, M., Schein, E., Rinder, H., Gothe, R., 1998, Characteristic genotypes discriminate

between Babesia canis isolates of differing vector specificity and pathogenicity to dogs.

Parasitol Res 84, 544-548.

Zinkl, J.G., Mcdonald, S.E., Kier, A.B., Cippa, S.J., Small, P.J., 1981, Cytauxzoon-like

organisms in erythrocytes of 2 cheetahs. J Am Vet Med Assoc 179, 1261-1262.

49

CHAPTER 2

DISTRIBUTION OF CYTAUXZOON FELIS IN BOBCATS, LYNX RUFUS, FROM THIRTEEN

STATES1

1Shock, Barbara C., Staci M. Murphy, Laura L. Patton, Philip M. Shock, Colleen Olfenbuttel, Jeff Beringer, Suzanne Prange, Daniel M. Grove, Matt Peek, Jay Butfiloski, Daymond W. Hughes, J. Mitchell Lockhart, Sarah Bevins, Victor F. Nettles, Holly M. Brown, David S. Peterson, and Michael J. Yabsley. 2010. Submitted to Veterinary Parasitology.

50

Abstract Cytauxzoon felis, a protozoan parasite of wild and domestic felids, is the causative agent

of cytauxzoonosis in domestic and some exotic felids in the United States. Previous research has

identified the bobcat (Lynx rufus) as the natural reservoir for this parasite. C. felis has been

experimentally transmitted by two tick species, Dermacentor variabilis and Amblyomma

americanum, which have overlapping distributions throughout the southeastern United States.

Our objective of the current study was to determine the distribution and prevalence of C. felis in

free-ranging bobcat populations from 13 states including California, Colorado, Florida, Georgia,

Kansas, Kentucky, Missouri, North Carolina, North Dakota, Ohio, Oklahoma, South Carolina,

and West Virginia). These states were selected because D. variabilis is present in each state, but

A. americanum is only currently known to be present in a subset of these states. Blood or spleen

samples from 697 trapper harvested or road-killed bobcats were tested for C. felis infection by a

polymerase chain reaction (PCR) assay which targets the first ribosomal internal transcribed

spacer region (ITS-1). Significantly higher prevalences of C. felis were detected from Missouri

(79%, n=39), North Carolina (63%, n=8), Oklahoma (60%, n=20), South Carolina (57%, n=7),

Kentucky (55%, n=74), Florida (44%, n=45), and Kansas (27%, n=41) compared with Georgia

(9%, n=159), North Dakota (2.4%, n=124), Ohio (0%, n=19), and West Virginia (0%, n=37). In

addition to bobcats, we tested a limited number of samples (n=8) from other wild felids and only

one cougar (Puma concolor) from Louisiana was positive, which represents the first report in an

infected cougar outside the Florida panther population. These data also indicate that C. felis is

present in North Dakota where C. felis has not been reported in domestic cats. Based on linear

regression analysis, prevalence rates were significantly higher in states where there are

established populations of A. americanum, which supports recent data on the experimental

51

transmission of C. felis by A. americanum and the fact that domestic cat cases are temporally

associated with A. americanum activity. Collectively, these data confirm that bobcats are a

common reservoir for C. felis and that A. americanum likely is an epidemiologically important

vector.

Introduction

Piroplasms in the genus Cytauxzoon (Family Theileridae) are related to members of the

genera Theileria and Babesia, and Cytauxzoon spp. are increasingly reported in domestic and

wild felid species worldwide (Luaces et al., 2005; Criado-Fiarello et al., 2004; Pexioto et al.,

2005; Ketz-Riley et al., 2000). The genera Cytauxzoon is currently restricted to felid species due

to the schizogenous replication in mononuclear phagocytes (Nijhof et al., 2005). Other

Cytauxzoon sp. previously described in African ungulates have since been reclassified as

Theileria spp. (Nijhof et al., 2005). In the United States, only one species, Cytauxzoon felis, has

been detected and it is an emerging infectious pathogen of domestic cats in Southeastern,

Midwestern, and Mid-Atlantic States (Wagner, 1976; Ferris, 1979; Kier et al., 1982b;

Birkenheuer et al., 2006). Cytauxzoon spp. are vectored by Ixodid ticks, and C. felis has been

experimentally transmitted by two ticks, Dermacentor variabilis and Amblyomma americanum

(Blouin et al., 1984; Kocan et al., 1992; Reichard et al., 2009; Edwards et al., 2010).

C. felis was first described in domestic cats from Missouri in 1976 (Wagner, 1976). Since

that time, C. felis has been detected in domestic cats (Felis catus) from numerous states

including Alabama, Arkansas, Florida, Georgia, Kansas, Kentucky, Louisiana, Mississippi,

Missouri, North Carolina, Oklahoma, South Carolina, Tennessee, Texas, and Virginia (Bendele

et al., 1976; Wagner, 1976, Wightman et al., 1977; Ferris, 1979; Glenn and Stair, 1982; Hauck,

52

1982; Kocan and Kocan, 1991; Meier and Moore, 2000; Birkenheuer et al., 2006; Haber et al.,

2007). Historically, infection with the parasite was considered uniformly fatal for domestic cats

due to the development of acute clinical cytauxzoonosis (Ferris, 1979). The pathognomonic sign

of C. felis infection in the domestic cat, occlusion of blood vessels by schizont-laden

macrophages, is also presumed to be responsible for much of the observed morbidity and

mortality. Recently, however, research and surveillance studies have indicated that some

domestic cats can survive infection and become persistently parasitemic (Kier et al., 1982b;

Meinkoth and Kocan, 2005; Haber et al., 2007; Brown et al., 2008).

The bobcat (Lynx rufus) is considered to be the natural reservoir for C. felis in the United

States (Kier et al., 1982a; Kier et al., 1982b; Glenn et al., 1983; Glenn and Stair; Blouin et al.,

1984). Naturally infected bobcats rarely display clinical signs and there is only one report from

Kansas of a naturally infected young bobcat dying of clinical cytauxzoonosis (Nietfeld and

Pollock, 2002). Experimental studies have shown that bobcats can develop acute cytauxzoonosis

when inoculated with schizogenous stages of the parasite, but no clinical signs were observed

when the bobcats were inoculated with the intraerythrocytic stages (Kier et al., 1982b; Glenn et

al., 1983) Similarly, bobcats experimentally infected via tick transmission typically have a

limited schizogenous phase which leads to a long-term subclinical parasitemia (Blouin et al.

1987). Few studies have examined the prevalence of the parasite within bobcat population, but

previously, high prevalences were detected in Oklahoma (31%-60%) and North Carolina (33%)

and a low prevalence was detected in Pennsylvania (7%) (Glenn et al., 1982; Glenn et al., 1983;

Kocan and Blouin, 1985; Birkenheuer et al., 2008).

C. felis has also infected other felids in Florida including free-ranging panthers (Puma

concolor) and a captive white tiger (Panthera tigris) (Butt et al., 1991; Rotstein et al., 1999;

53

Garner et al., 1996). The tiger died of acute cytauxzoonosis. In the endangered Florida panther

populations, C. felis has been described as causing mild hemolytic anemia and liver damage, but

no deaths have been attributed to the parasite (Yabsley et al., 2006; Harvey et al., 2007). Free-

ranging cougars may therefore be another natural, subclinical reservoir for C. felis.

In the current study, we conducted a comprehensive study of the distribution and

prevalence of C. felis in free-ranging bobcat populations from thirteen states. Limited cougar

samples from three other states were included in the study as well as one serval sample

(Leptailurus serval). The objective of this project was to gain a better understanding of the

natural history of the parasite within wild felid populations with the hope of increasing the health

of domestic, exotic, and wild felids in the United States.

Materials and Methods

Sample collection

Samples were collected opportunistically from a variety of sources including trapper-

harvested, road-killed, or clinical case submissions of mortalities. Spleen samples were either

collected from fresh carcasses and frozen at -20 C prior to DNA extraction or entire carcasses

were frozen and later necropsied to collect spleen samples. When fresh carcasses were available,

a blood sample was collected from the heart or thoracic cavity. From 1999 to 2010, 697 blood or

spleen samples were collected from as many bobcats from thirteen states (California, Colorado,

Florida, Georgia, Kansas, Kentucky, Missouri, North Carolina, North Dakota, Ohio, Oklahoma,

South Carolina, and West Virginia; Figure 1). Additionally, seven cougar samples were collected

from Georgia (n=1), Louisiana (n=1), and North Dakota (n=5) and one escaped exotic serval

(genus species) sample was collected from Louisiana.

54

Molecular Analysis

Genomic DNA was extracted from100μl of whole blood or 10 mg spleen using the

Qiagen DNA Purification Kit (Germantown, MD) following the manufacturer’s protocol. A

nested PCR protocol was used that amplifies the entire internal transcribed spacer (ITS)-1 rRNA

region of most piroplasms including Cytauxzoon, Babesia, and Theileria spp. (Bostrom et al.,

2008). Briefly, for primary amplification, 5 μl of DNA was added to 20 μl of a master mix

containing 10 mM Tris-Cl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.2 mM each dNTP (Promega,

Madison, Wisconsin), 2.5 units Taq DNA Polymerase (Promega), and 0.8 μM of primers ITS-

15C (5’-CGATCGAGTGATCCGGTGAATTA) and ITS-13B (5’-

GCTGCGTCCTTCATCGTTGTG). Cycling parameters were 94 C for 1 min followed by 35

cycles of 94 C for 30 sec, 52 C for 30 sec, 72 C for 1 min, and a final extension at 72 C for 5

min. For the nested PCR, 1 μl of primary product was used as template in a 25 μl reaction

containing the same PCR components except primers, ITS-15D (5’-

AAGGAAGGAGAAGTCGTAACAAGG) and ITS-13C (5’-

TTGTGTGAGCCAAGACATCCA) were used. The cycling parameters were the same as the

primary reaction except the annealing temperature was 49 C.

To prevent and detect contamination, DNA extraction, primary and secondary

amplification, and product analysis were done in separate dedicated areas. A negative water

control was included in each set of DNA extraction, and a different water control was included in

each set of primary and secondary PCR reactions. All amplicons of approximately 550 bp were

purified with a Qiagen gel extraction kit (Germantown, MD) and bi-directionally sequenced at

the University of Georgia Integrated Biotechnology Laboratory (Athens, GA).

55

Data Analysis

Linear regression and chi-square analysis were performed using MiniTab version 16

statistical software.

Results

Based on PCR testing and sequence analysis, 138 of the 697 (20%) bobcats were positive

for C. felis (Table 1). and infected bobcats were detected in all states except California,

Colorado, Ohio, and West Virginia. Significantly higher prevalence rates (>50%) were detected

in Kentucky (55%), South Carolina (57%), North Carolina (63%), Oklahoma (65%), and

Missouri (79%) (Table 2.1). All amplicons were sequenced and confirmed to be C. felis. A

limited number of bobcats had positive amplification with our PCR protocol but were

determined to be infected with other protozoan parasites (e.g. Heptazoon spp., Toxoplasma

gondii, and Babesia spp.).

All states in this study have reported the presence of D. variabilis, but A. americanum has

not been reported from California, Colorado, or North Dakota and only low prevalences have

been reported from southern Ohio and West Virginia. Simple linear regression (y= 0.004 +

0.49x) showed that a high prevalence of A. americanum in a state was significantly correlated

with a higher prevalence of C. felis in bobcats (F= 22.89, p= 0.001). No sex differences in C.

felis infection rate were seen in the bobcat populations. Chi-squre analysis revealed that there

were significant differences in prevalences between states (Table 2.1). Although limited

numbers of other free-ranging felids were tested, we detected C. felis in a single panther from

Louisiana.

56

Discussion

This is the most comprehensive study of the distribution and prevalence of C. felis in

bobcats, the natural wildlife reservoir of C. felis in the United States. A total of 707 felids from

fourteen states were examined for the parasite. Because C. felis prevalence is higher in wild

felids compared with domestic cats, testing of wild felids provides a more sensitive method to

determine the distribution and prevalence of C. felis in the United States. Interestingly, we

detected a low prevalence of C. felis in North Dakota which is the first report of C. felis in any

felid from this state as well as the most northerly report of C. felis since Birkenheuer et al. (2008)

reported the parasite in free-ranging bobcats from Pennsylvania. In addition, we detected a C.

felis positive cougar in Louisiana which represents the first report of C. felis in a cougar outside

the state of Florida. Although C. felis has been reported in domestic cats from Georgia,

Kentucky, and South Carolina, this is the first report of C. felis from a free-ranging wild felid in

these states.

We found a significant correlation between higher prevalence rates of C. felis in states

where both confirmed tick vectors are known to be present and common. These data are in

agreement with reports of clinical cytauxzoonosis in domestic cats which only been reported

from Southeastern, Midwestern, and Mid-Atlantic states, but not from California, Colorado,

North Dakota, Ohio, or West Virginia. Of all the states where A. americanum numbers are low to

absent, C. felis was only reported in North Dakota (2%). North Dakota is well outside the known

range of A. americanum, which is not reported to be north of Iowa or west of Nebraska (CDC,

2009), although unpublished reports of A. americanum exist for South Dakota (M. Wimberly,

unpublished data). Aside from the prevalence of the state of Georgia (9%), all other prevalences

57

were above 30%. These field data suggest that A. americanum may play a more primary role in

the maintenance and dissemination of the parasite in wild felids. Recent tick transmission studies

have indicated that A. americanum may be a more competent vector of C. felis than D. variabilis

as in repeated trials the latter was an unsuccessful vector whereas A. americanum transstadially

transmitted the parasite under identical situations (Reichard et al., 2009; Edwards et al., 2010).

Additionally, C. felis has only been detected from wild-caught questing A. americanum from

Oklahoma (MIR 0.5-1.5%), while there is no report of C. felis from questing D. variabilis

(Edwards et al., 2010).

Most of the information about the about the status of L. rufus as a reservoir for C. felis

was determined by studies on Oklahoma bobcats in the 1980s (Kier et al., 1982a; Kier et al.,

1982b; Kocan and Blouin, 1985; Blouin et al., 1987) which revealed high prevalences of

subclinical infection with C. felis (31-60%). A study by Birkenheuer et al. (2008) also revealed a

prevalence of C. felis infection in bobcats from North Carolina (33%), which the data from this

study supports (63%). This study indicates that C. felis is more widespread and prevalent in

bobcat populations than previously suspected. The high prevalences observed in presumably

healthy free-ranging wild felids are further confirmation of L. rufus as the primary reservoir for

the parasite. While no felids in this study died of clinical cytauxzoonosis, experimental animals

and the of naturally infected young bobcat dying of cytauxzoonosis suggest that some bobcats

die of C. felis each year, undetected. Molecular characterization of C. felis detected in domestic

cats from Arkansas and Georgia suggest that there are strains of C. felis circulating in domestic

cat which varying in their pathogenicity (Brown et al., 2009a; Brown et al., 2009b). Domestic

cats which become persistently parasitemic can in turn become a reservoir for wild felids.

58

Acknowledgements The authors thank numerous personnel from state agencies who collected felid samples. This

study was primarily funded by the Morris Animal Foundation (DO8FE-003). Additional support

was provided by the Federal Aid to Wildlife Restoration Act (50 Stat. 917) and through

sponsorship from fish and wildlife agencies in Alabama, Arkansas, Florida, Georgia, Kansas,

Kentucky, Louisiana, Maryland, Mississippi, Missouri, North Carolina, Oklahoma, Puerto Rico,

South Carolina, Tennessee, Virginia, and West Virginia.

References Bendele, R.A., Schwartz, W.L., Jones, L.P.,1976, Cytauxzoon-like disease in Texas cats.

Southwest Vet 29, 244-246.

Birkenheuer, A.J., Marr, H., Alleman, A.R., Levy, M.G., Breitschwerdt, E.B., 2006,

Development and evaluation of a PCR assay for the detection of Cytauxzoon felis DNA

in feline blood samples. Vet Parasitol 137, 144-149.

Birkenheuer, A.J., Marr, H.S., Warren, C., Acton, A.E., Mucker, E.M., Humphreys, J.G., Tucker,

M.D., 2008, Cytauxzoon felis infections are present in bobcats (Lynx rufus) in a region

where cytauxzoonosis is not recognized in domestic cats. Vet Parasitol 153, 126-130.

Blouin, E.F., Kocan, A.A., Glenn, B.L., Kocan, K.M., Hair, J.A., 1984, Transmission of

Cytauxzoon felis Kier, 1979 from bobcats, Felis rufus (Schreber), to domestic cats by

Dermacentor variabilis (Say). J Wildl Dis 20, 241-242.

Blouin, E.F., Kocan, A.A., Kocan, K.M., Hair, J., 1987, Evidence of a limited schizogonous

cycle for Cytauxzoon felis in bobcats following exposure to infected ticks. J Wildlife Dis

23, 499-501.

59

Bostrom, B., Wolf, C., Greene, C., Peterson, D.S., 2008, Sequence conservation in the rRNA

first internal transcribed spacer region of Babesia gibsoni genotype Asia isolates. Vet

Parasitol 152, 152-157.

Brown, H.M., Berghaus, R.D., Latimer, K.S., Britt, J.O., Rakich, P.M., Peterson, D.S., 2009,

Genetic variability of Cytauxzoon felis from 88 infected domestic cats in Arkansas and

Georgia. J Vet Diagn Invest 21, 59-63.

Brown, H.M., Modaresi, S.M., Cook, J.L., Latimer, K.S., Peterson, D.S., 2009, Genetic

variability of archived Cytauxzoon felis histologic specimens from domestic cats in

Georgia, 1995-2007. J Vet Diagn Invest 21, 493-498.

Brown, H.M., Latimer, K.S., Erikson, L.E., Cashwell, M.E., Britt, J.O., Peterson, D.S., 2008,

Detection of persistent Cytauxzoon felis infection by polymerase chain reaction in three

asymptomatic domestic cats. J Vet Diagn Invest 20, 485-488.

Butt, M.T., Bowman, D., Barr, M.C., Roelke, M.E., 1991, Iatrogenic transmission of Cytauxzoon

felis from a Florida panther (Felis concolor coryi) to a domestic cat. J Wildl Dis 27, 342-

347.

Criado-Fornelio, A., Gonzalez-del-Rio, M.A., Buling-Sarana, A., Barba-Carretero, J.C., 2004,

The "expanding universe" of piroplasms. Vet Parasitol 119, 337-345.

Edwards, A.C., Reichard, M.V., Meinkoth, J.H., Snider, T.A., Meinkoth, K.R., Heinz, R.E.,

Little, S.E., 2010, Confirmation of Amblyomma americanum (Acari: Ixodidae) as a vector

for Cytauxzoon felis (Piroplasmorida: Theileriidae) to domestic cats. J Med Ent.

Ferris, D.H., 1979, A progress report on the status of a new disease of American cats:

cytauxzoonosis. Comp Immunol Microbiol Infect Dis 1, 269-276.

60

Garner, M.M., Lung, N.P., Citino, S., Greiner, E.C., Harvey, J.W., Homer, B.L., 1996, Fatal

cytauxzoonosis in a captive-reared white tiger (Panthera tigris). Vet Pathol 33, 82-86.

Glenn, B.L., Kocan, A.A., Blouin, E.F., 1983, Cytauxzoonosis in bobcats. J Am Vet Med Assoc

183, 1155-1158.

Glenn, B.L., Rolley, R.E., Kocan, A.A., 1982, Cytauxzoon-like piroplasms in erythrocytes of

wild-trapped bobcats in Oklahoma. J Am Vet Med Assoc 181, 1251-1253.

Haber, M.D., Tucker, M.D., Marr, H.S., Levy, J.K., Burgess, J., Lappin, M.R., Birkenheuer, A.J.,

2007, The detection of Cytauxzoon felis in apparently healthy free-roaming cats in the

USA. Vet Parasitol 146, 316-320.

Hauck, W.N., Snider, T.G., Lawrence, J.E., 1982, Cytauxzoonosis in a native Louisiana cat, J

Am Vet Med Asso 180, 1472-1474.

Harvey, J.W., Dunbar, M.R., Norton, T.M., Yabsley, M.J., 2007, Laboratory findings in acute

Cytauxzoon felis infection in cougars (Puma concolor couguar) in Florida. J Zoo Wildlife

Med 38, 285-291.

Ketz-Riley, C.J., Reichard, M.V., Van Den Bussche, R.A., Hoover, J.P., Meinkoth, J., Kocan,

A.A., 2003, An intraerythrocytic small piroplasm in wild-caught Pallas's cats

(Otocolobus manul) from Mongolia. J Wildlife Dis 39, 424-430.

Kier, A.B., Wagner, J.E., Morehouse, L.G., 1982a, Experimental Transmission of Cytauxzoon

felis from Bobcats (Lynx rufus) to Domestic Cats (Felis domesticus). Am J Vet Res 43,

97-101.

Kier, A.B., Wightman, S.R., Wagner, J.E., 1982b, Interspecies Transmission of Cytauxzoon felis.

Am J Vet Res 43, 102-105.

61

Kocan, A.A., Kocan, K.M., 1991, Tick-transmitted diseases of wildlife in North America. Bull

Soc Vector Eco 16, 94-108.

Kocan, A.A., Kocan, K.M., Blouin, E.F., Mukolwe, S.W., 1992, A Redescription of Schizogony

of Cytauxzoon felis in the Domestic Cat. Ann Ny Acad Sci 653, 161-167.

Luaces, I., Aguirre, E., Garcia-Montijano, M., Velarde, J., Tesouro, M.A., Sanchez, C., Galka,

M., Fernandez, P., Sainz, A., 2005, First report of an intraerythrocytic small piroplasm in

wild Iberian lynx (Lynx pardinus). J Wildlife Dis 41, 810-815.

Meier, H.T., Moore, L.E., 2000, Feline cytauxzoonosis: A case report and literature review. J

Am Anim Hosp Assoc 36, 493-496.

Meinkoth, J.H., Kocan, A.A., 2005, Feline cytauxzoonosis. Vet Clin N Am-Small 35, 89-+.

Nietfeld, J.C., Pollock, C., 2002, Fatal cytauxzoonosis in a free-ranging bobcat (Lynx rufus). J

Wildlife Dis 38, 607-610.

Nijhof, A.M., Pillay, V., Steyl, J., Prozesky, L., Stoltsz, W.H., Lawrence, J.A., Penzhorn, B.L.,

Jongejan, F., 2005, Molecular characterization of Theileria species associated with

mortality in four species of African antelopes. J Clin Microbiol 43, 5907-5911.

Peixoto, P.V., Soares, C.O., Scofield, A., Santiago, C.D., Franca, T.N., Barros, S.S., 2007, Fatal

cytauxzoonosis in captive-reared lions in Brazil. Vet Parasitol 145, 383-387.

Reichard, M.V., Meinkoth, J.H., Edwards, A.C., Snider, T.A., Kocan, K.M., Blouin, E.F., Little,

S.E., 2008, Transmission of Cytauxzoon felis to a domestic cat by Amblyomma

americanum. Vet Parasitol.

Rotstein, D.S., Taylor, S.K., Harvey, J.W., Bean, J., 1999, Hematologic effects of

cytauxzoonosis in Florida panthers and Texas cougars in Florida. J Wildlife Dis 35, 613-

617.

62

Wagner, J.E., 1976, A fatal cytauxzoonosis-like disease in cats. J Am Vet Med Assoc 168, 585-

588.

Wightman, S.R., Kier, A.B., Wagner, J.E., 1977, Feline cytauxzoonosis: clinical features of a

newly described blood parasite disease. Feline Pract, 24-26.

Yabsley, M.J., Murphy, S.M., Cunningham, M.W., 2006, Molecular detection and

characterization of Cytauxzoon felis and a Babesia species in cougars from Florida. J

Wildlife Dis 42, 366-374.

63

Table 2.1. Prevalence of Cytauxzoon felis in bobcats. Percentages with different letters are

significantly different (p<0.05).

State Tick species prevalent No.positive for C. felis/No.tested

(% infected)

California Dv 0/26 (0%) a

Colorado Dv 0/67 (0%) a

Ohio Dv 0/19 (0%) a

West Virginia Dv 0/37 (0%) a

North Dakota Dv 3/172 (2%) a

Georgia Dv and Aa 13/143 (9%) b

Kansas Dv and Aa 12/39 (31%) c

Florida Dv and Aa 16/45 (36%) c

Kentucky Dv and Aa 41/74 (55%) c

South Carolina Dv and Aa 4/7 (57%) c

North Carolina Dv and Aa 5/8 (63%) c

Oklahoma Dv and Aa 13/20 (65%) c, d

Missouri Dv and Aa 31/39 (79%) d

138/697

64

Figure 2.1. Distribution of Cytauxzoon felis in bobcats. Light grey counties represent counties

from which bobcats and/or cougars were sampled. Dark grey counties that at least one positive

wild felid was detected.

65

CHAPTER 3

EXTENSIVE GENETIC VARIABILITY OF CYTAUXZOON FELIS FROM BOBCATS (LYNX

RUFUS) AND COUGARS (PUMA CONCOLOR)

1Shock, Barbara C., Staci M. Murphy, Laura L. Patton, Philip M. Shock, Colleen Olfenbuttel, Jeff Beringer, Suzanne Prange, Daniel M. Grove, Matt Peek, Jay Butfiloski, Daymond W. Hughes, J. Mitchell Lockhart, Victor F. Nettles, Holly M. Brown, Adam J. Birkenheuer, David S. Peterson, and Michael J. Yabsley. 2010. To be submitted to Veterinary Parasitology.

66

Abstract

Cytauxzoon felis, a tick-borne protozoan parasite, is the causative agent of

cytauxzoonosis in domestic cats in the United States. The natural reservoir for this parasite is the

bobcat, Lynx rufus, which typically develops only limited or no clinical signs of illness.

Although not likely important reservoirs, C. felis has also been detected in free-ranging Florida

panthers (Puma concolor coryii). Recent studies have identified specific genotypes of C. felis

circulating in domestic cats which are associated with different clinical outcomes and specific

spatial locations. In the current study, we investigated the intraspecific variation of the C. felis

internal transcribed spacer (ITS)-1 and ITS-2 rRNA regions from 153 bobcats and eight cougars

from 11 states (Florida, Georgia, Kansas, Kentucky, Louisiana, Missouri, North Carolina, North

Dakota, South Carolina, Oklahoma, and Pennsylvania). Unambiguous ITS-1 and ITS-2 data

were obtained for 131 and 106 samples, respectively, and both ITS-1 and ITS-2 sequences were

obtained for 82 (51%) samples. Within the ITS-1 region, sequences from 46 bobcats were

identified which were identical to those previously reported in domestic cats and we identified 60

unique sequences. Samples from 46 bobcats and one cougar had ITS-1 sequences identical to the

most common sequence reported from domestic cats. The most common ITS-2 sequence from

domestic cats was also common in wild felids (36 bobcats and a cougar). Five other ITS-2

sequences had been previously reported in domestic cats; the remaining ITS-1 and ITS-2

sequences from wild felids were novel. Samples from three Florida panthers and a bobcat from

MO had a 40/41-bp insert in the ITS-2 similar to one described previously in a cat from AR.

Additionally, a previously undescribed 198/199-bp insert was detected in the ITS-2 sequence

from in four bobcats. Collectively, nine bobcats and one cougar were infected with genotype

ITSA and two bobcats were infected with genotypes ITSB and ITSC. Eight bobcats from

67

southwestern Georgia and northwestern Florida were infected with a novel genotype (ITSD).

These data indicate that based on ITS1 and ITS2 sequences, numerous C. felis strains circulate in

wild felids and that some genotypes may cluster in specific spatial locations.

Introduction

Cytauxzoon felis, a tick-borne protozoan parasite, is the causative agent of

cytauxzoonosis in domestic cats in the United States. The parasite was first identified in

domestic cats from Missouri, Texas and Arkansas in the 1970s (Bendele et al., 1976; Wagner,

1976; Wightman et al., 1977) and has subsequently been identified in domestic cats from

numerous southeastern, Midwestern, and Mid-Atlantic states (Wagner, 1976; Ferris, 1979; Kier

et al., 1982b; Birkenheuer et al., 2006). In domestic cats, C. felis infection causes erythrocyte

hemolysis and occlusion of the lumen of blood vessels by large schizont-laden mononuclear

phagocytes in the lungs, liver, lymph nodes, and spleen (Simpson et al., 1985; Kier et al., 1987;

Kocan and Kocan, 1991; Kocan et al., 1992). Historically, the mortality rate due to this parasite

was near100%; however, recent studies have discovered an increasing number of domestic cats

that have subclinical chronic infections (Ferris, 1979; Haber et al., 2007; Birkenheuer et al.,

2006). Both Amblyomma americanum (lone star tick) and Dermacentor variabilis (American dog

tick) are confirmed vectors. Both tick species are common in the southeastern and Midwestern

United states where the majority of C. felis infections have been identified (Blouin et al., 1984;

Kocan et al., 1992; Reichard et al., 2009; Edwards et al., 2010; Shock et al., submitted).

The bobcat (Lynx rufus) is considered to be the primary wildlife reservoir. High

prevalences of C. felis infections have been documented in bobcat populations (Lynx rufus) from

Missouri, Oklahoma, and North Carolina (Wagner, 1976; Glenn et al., 1983; Birkenheuer et al.,

68

2008). Experimental and field-based studies indicate that the majority of infected bobcats suffer

no clinical disease; however, rare cases of experimental and natural infections can result in

mortality (Kier et al., 1982a; Kier et al., 1982b; Glenn et al., 1983; Blouin et al., 1984; Blouin et

al., 1987; Neitfeld and Pollock, 2002). For example, a bobcat experimentally inoculated with

schizogenous stages of the parasite died of acute cytauxzoonosis, while bobcats experimentally

infected via tick transmission only developed a limited schizogenous phase which lead to long-

term subclinical parasitemia (Kier et al., 1982a; Blouin et al. 1987). A single report of mortality

of a young bobcat from C. felis infection has been reported in Kansas (Nietfeld and Pollock,

2002). Florida panthers (Puma concolor coryii) have also been identified as a possible reservoir

for C. felis, although the parasite appears to cause a mild hemolytic anemia (Butt et al., 1991;

Yabsley et al., 2006; Harvey et al. 2007). Worldwide, Cytauxzoon spp. that are distinct from C.

felis, have been identified in numerous wild felid species (Luaces et al., 2005; Pexioto, 2005;

Ketz-Riley et al., 2003; Butt et al., 1991).

Increasing reports of subclinical infections that appear to be unrelated to treatment

(Birkenheuer et al,. 2006; Haber et al., 2007; Brown et al., 2008) suggest that different strains of

C. felis may differ in their virulence for domestic cats or that cats are evolving resistance to

clinical disease associated with this parasite. Currently there is no data on virulence genes for C.

felis, but two recent studies have examined genetic variation within the internal transcribed

spacer (ITS) regions of the ribosomal RNA genes (Brown et al., 2009a; Brown et al., 2009b).

These two studies identified several genotypes and numerous unique gene sequences in C. felis

samples from domestic cats from Georgia and Arkansas. Significant spatial correlations and

associations with clinical outcome were associated with specific genotypes (Brown et al.,

2009a). These data suggest that specific genotypes are associated with clinical outcome in

69

domestic cats, and if confirmed, these genetic markers could be used to better predict the clinical

outcome and to guide treatment and management protocols (Brown et al., 2009a). Currently,

there is no information regarding the genetic variation among C. felis samples from wild felids in

the United States. We hypothesize that testing of wild felid samples will reveal additional

genotypes and that spatial correlations with genotype will be more easily identified because of

the high prevalence of C. felis these reservoirs and the lack of movement of wild felids to new

geographic regions.

Materials and Methods

Samples

We included DNA samples from 161 Cytauxzoon felis-infected bobcats (n=153) and

cougars (n=8) that were determined to be positive during three previous studies on these wild

felids (Yabsley et al., 2006; Birkenheuer et al., 2008; Shock et al., unpublished). These samples

were confirmed positive for C. felis by either amplification of a segment of the 18S rRNA gene

or the ITS-1 region followed by sequence analysis. All DNA samples were held at -20C or -80C

until testing in the current study.

Genetic characterization

Two different regions were targeted to identify genetic variability, the ITS-1 and ITS-2

regions. The ITS-1 region was amplified using a nested PCR that amplifies this genetic region

from most piroplasms including Cytauxzoon, Babesia, and Theileria spp. (Bostrom et al., 2008).

Briefly, for primary amplification, 5 μl of DNA was added to 20 μl of a master mix containing

10 mM Tris-Cl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.2 mM each dNTP (Promega, Madison,

Wisconsin), 2.5 units Taq DNA Polymerase (Promega), and 0.8 μM of primers ITS-15C (5’-

70

CGATCGAGTGATCCGGTGAATTA) and ITS-13B (5’-GCTGCGTCCTTCATCGTTGTG).

Cycling parameters were 94 C for 1 min followed by 35 cycles of 94 C for 30 sec, 52 C for 30

sec, 72 C for 1 min, and a final extension at 72 C for 5 min. For the nested PCR, 1 μl of primary

product was used as template in a 25 μl reaction containing the same PCR components except

primers ITS-15D (5’-AAGGAAGGAGAAGTCGTAACAAGG) and ITS-13C (5’-

TTGTGTGAGCCAAGACATCCA) were used. The cycling parameters were the same as the

primary reaction except the annealing temperature was 49 C. A single PCR was used to amplify

the ITS-2 region. The same master mix protocol was used except for the inclusion of primers

FOR7 (5’ -AGCCAATTGCGATAAGCATT) and REV7 (5’-

TCACTCGCCGTTACTAGGAGA) and the cycling parameters were 96 for 3 min followed by

30 cycles of 94 C for 30 sec, 60 C for 30 sec, 72 C for 1 min 30 sec, and a final extension at 72 C

for 7 min.

To prevent and detect contamination, primary and secondary amplification, and product

analysis were done in separate dedicated areas. A negative water control was included in each set

of DNA extraction, and one water control was included in each set of primary and secondary

PCR reactions. All amplicons of the appropriate size (~550 bp for ITS1 and ~300 bp for ITS2)

were purified with a Qiagen gel extraction kit (Germantown, MD) and bi-directionally

sequenced at the University of Georgia Integrated Biotechnology Laboratory (Athens, GA).

Chromatogram data were analyzed using Sequencher (Ann Arbor, MI) and sequences

with ambiguous bases (other than simple incorporation of two nucleotides at a single position)

were excluded from the study. Sequences obtained from this study and available in GenBank

were aligned using the multisequence alignment ClustalX program within MEGA (Molecular

Evolutionary Genetics Analysis) version 3.1 program (Kumar et al., 2004).

71

Results

Amplification and sequencing of the two ITS regions of C. felis from samples previously

determined to be infected with C. felis yielded unambiguous ITS1 data for 131 of 161 (81%)

samples from GA, FL, KS, KY, LA, MO, NC, ND, OK, PA and SC and ITS2 data for 106 of

161 (66%) samples from GA, FL, KY, LA, MO, NC, ND, OK, PA and SC. Combined ITS1 and

ITS2 sequence data was available for 82 of the 161 samples (51%). Assignment of samples to a

particular genotype was only conducted on samples that had both ITS1 and ITS2 data (as

described by Brown et al., 2009a, 2009b) which included felids from Georgia (n=7 bobcats),

Florida (n=12 bobcats, 1 Florida panther), Kentucky (n=27 bobcats), Missouri (n=22 bobcats),

North Carolina (n=6 bobcats), Oklahoma (n=6 bobcats), and Louisiana (1 cougar).

Within the 458-bp ITS1 region, there were 100 single nucleotide polymorphisms (SNPs)

or insertions/deletions. The most common ITS-1 sequence detected in this study was identical to

EU450802 and was found in 46 bobcats (2 from GA, 2 from FL, 21 from KY, 3 from KS, 9 from

MO, 2 from OK, and 7 from NC) and in the cougar from LA. The second most common ITS-1

sequence was detected in 18 bobcats from FL (n=12) and GA (n=4). Two sequences from OK,

one each from MO and NC, and one each from KS and NC were identical to each other. The

remaining 60 sequences were unique.

Within the 265-bp ITS2 region of C. felis (excluding the two insertions mentioned

below), there was a total of 184 SNPs and 10 single nucleotide insertions, and 12 single

nucleotide deletions. Sequences from 36 bobcats (one each from GA, NC, and ND, three from

FL, 5 from OK, 12 from KY, and 13 from MO) and the cougar from LA had identical sequences

to one previously detected in domestic cats in AR and GA (EU450804). Sequences from single

bobcats from GA and SC were identical to EU450805 and FJ536418, respectively. Three bobcats

72

from MO, GA, and KY had sequences identical to FJ536419. Additional sequences that were

identical to each other were identified in multiple bobcats including nine samples from GA and

FL, four samples from KY, three samples from FL, two samples from KY, and two samples from

NC and MO.

Three ITS-2 sequences from Florida panthers had a 40bp insert nearly identical (38 or 39

of 40 bases) to an insert in a sequence from a single domestic cat (EU450806) from AR;

however, numerous SNPs within the non-insert region of the ITS2 distinguished these sequences

from EU450806. An additional sample from a bobcat from MO had a 41bp insert at the sample

location as EU450806 and the samples from the Florida panthers. Two samples from bobcats

from NC had a 199bp insert after position 152 (EU450804) that were 94.4% identical to each

other. Two additional bobcats from PA and KY had a similar 198bp insert (98.9% identical to

each other and 94.9-95.9% similar to the NC samples).

When the ITS1 and ITS2 sequences were combined, ten wild felids (nine bobcats from

FL (1), KY (2), OK (2), and MO (4) and the cougar from LA) were infected with the C. felis

ITSA genotype that is common in domestic cats (Brown et al., 2009a),. Genotype ITSB (from

Brown et al., 2009a) and sequence type ITSd (from Brown et al., 2009b) were each detected in a

single bobcat from GA. Based on the criteria of Brown et al., (2009a) we identified a novel

genotype (ITSD) from eight bobcats from GA (2) and FL (6).

Discussion

In the current study, we characterized the genetic variation in the ITS1 and ITS2 regions

of C. felis from wild felids, primarily bobcats, from numerous states in the eastern and central

United States. Compared with two previous studies (Brown et al., 2009a; Brown et al., 2009b)

73

and a limited number of other sequences available in GenBank, we observed more

polymorphisms in both of these genetic targets including SNPs and individual

insertions/deletions in both ITS1 and ITS2 and a novel 198/199bp insert in the ITS2. This

increased genetic diversity is not surprising because bobcats are the natural reservoirs of C. felis

and infections have been documented as early as 1930 (Wenyon and Hamerton, 1930). In

addition to being the primary reservoir, bobcats are useful for this type of project because they

are exposed to more potentially infected ticks than domestic cats and prevalence rates in bobcat

populations in some regions may exceed 50% which provides greater numbers of parasites for

characterization at single locations (Shock et. al., submitted). Unfortunately, many bobcats in the

current project appeared to be coinfected with multiple strains which created difficulty in

sequence analysis. Also, infected bobcats have been identified in regions where domestic cat

infections have not been reported (Birkenheuer et al., 2008; Shock et al., submitted).

Recently there have been increasing reports of natural subclinical chronically-infected

domestic cats as was normally only seen in wild felids infected with C. felis (Birkenheuer et al.,

2006; Haber et al., 2007; Brown et al., 2008). There have been at least three reasons postulated

for the increased recognition of chronically infected asymptomatic cats: 1) better treatment

strategies, 2) better diagnostics, or 3) strains of C. felis that differ in their virulence for domestic

cats. Although many treatment strategies have been attempted in reducing the parasitemia in

domestic cats (Green et al., 1999; Motzel and Wagner, 1995), none have been consistently

effective. Although diagnostics, particularly molecular techniques, have improved, previous

experimental studies suggested that different strains of C. felis had variable pathogenicity for cats

as some strains induced clinical disease while others induced subclinical chronic infections (Kier

et al., 1982b).

74

The first study to genetically characterize a large number of C. felis samples was

conducted on samples from domestic cats from AR and GA (Brown et al., 2009a). Significantly,

one genotype (ITSA) of combined ITS1 and ITS2 sequences was associated with an increased

survival rate of the infected domestic cats while cats infected with the two other genotypes

(ITSB and ITSC) were more likely to die from the infection. Interestingly, a possible geographic

association was found in the study; a significant proportion (84%) of the C. felis samples from

AR were identified as the ITSA genotype, and the majority of samples (68%) from GA were

classified as genotype ITSB (Brown et al., 2009a). However, a subsequent study of only GA

samples found that ITSC was the predominate genotype followed by genotype ITSB (Brown et

al., 2009b). Genotypes ITSA and ITSB can only be differentiated based on ITS-2 sequence

because they have identical ITS-1 sequences. In the current study, we detected the ITSA

genotype in nine bobcats from multiple states (FL, KY, OK, and MO) and the single cougar from

LA, but the majority of positives were from the central US (90%). Unfortunately, no bobcats

from AR were available for testing. The ITSB genotype was found in a single bobcat from GA,

which in one study was the predominate genotype detected in domestic cats from Georgia

(Brown et al., 2009a). Interestingly, the ITS-1 sequence (EU450802) that was most common in

both previous studies was the most common ITS-1 sequence detected in wild felids. Similarly,

the individual ITS-2 sequence that was most common in wild felids was the most commonly

found sequence in domestic cats (EU45804)

No significant geographic clustering was observed for the individual ITS1 and ITS2

sequences which were in general, found throughout the study area, including the most common

ITS1 and ITS2 sequences which were found in 46 bobcats and the cougar from LA. However,

we did identify a novel genotype (ITSD) in 8 bobcats that was restricted to northern FL and

75

southern GA (sampling sites in these two areas were very close to each other). The identification

of a novel genotype in this region was likely because a large number of samples were collected

from a small geographic region where no cat samples have previously been tested (Brown et al.,

2009a; Brown et al., 2009b). In the current study, our samples were opportunistically collected

which resulted in few individuals being sampled at the majority of our sites so although

numerous novel sequence types were detected, only one novel genotype was identified.

Alternatively, domestic cats may be more like to become infected with only certain genotypes or

genetic sequences (Brown et al., 2009a; Brown et al., 2009b). Additional samples from both wild

and domestic felids are needed from geographically discrete areas to detect if clustering of

sequence types or selection in domestic cats is occurring.

It is important to note that neither of the sequenced regions examined in the current

study, or previous ones (Brown et al., 2009a, 2009b), are related to virulence genes or any

potential protein involved in pathogenicity. They are simply being used as potential markers for

observed biological or spatial differences. Future studies should target genes that are associated

with pathogenicity or antigenic variation in related pathogens (e.g., Babesia spp.) such as the

ones that encode for the variant erythrocyte surface antigen-1 (VESP1), leucine aminopeptidase,

or, heat shock protein-70 proteins (Yamasaki et al., 2007; Jia et al., 2009; Xiao et al., 2010)

Acknowledgements

The authors thank numerous personnel from state agencies who collected felid samples.

This study was primarily funded by the Morris Animal Foundation (DO8FE-003). Additional

support was provided by the Federal Aid to Wildlife Restoration Act (50 Stat. 917) and through

SCWDS sponsorship from fish and wildlife agencies in Alabama, Arkansas, Florida, Georgia,

76

Kansas, Kentucky, Louisiana, Maryland, Mississippi, Missouri, North Carolina, Oklahoma,

Puerto Rico, South Carolina, Tennessee, Virginia, and West Virginia.

References

Bendele, R.A., Schwartz, W.L., Jones, L.P.,1976, Cytauxzoon-like disease in Texas cats.

Southwest Vet 29, 244-246.

Barta, J.R., Coles, B.A., Schito, M.L., Fernando, M.A., Martin, A., Danforth, H.D., 1998,

Analysis of infraspecific variation among five strains of Eimeria maxima from North

America. Int J Parasitol 28, 485-492.

Birkenheuer, A.J., Le, J.A., Valenzisi, A.M., Tucker, M.D., Levy, M.G., Breitschwerdt, E.B.,

2006, Cytauxzoon felis infection in cats in the mid-Atlantic states: 34 cases (1998-2004).

J Am Vet Med Assoc 228, 568-571.

Birkenheuer, A.J., Marr, H.S., Warren, C., Acton, A.E., Mucker, E.M., Humphreys, J.G., Tucker,

M.D., 2008, Cytauxzoon felis infections are present in bobcats (Lynx rufus) in a region

where cytauxzoonosis is not recognized in domestic cats. Vet Parasitol 153, 126-130.

Blouin, E.F., Kocan, A.A., Glenn, B.L., Kocan, K.M., Hair, J.A., 1984, Transmission of

Cytauxzoon felis Kier, 1979 from bobcats, Felis rufus (Schreber), to domestic cats by

Dermacentor variabilis (Say). J Wildl Dis 20, 241-242.

Blouin, E.F., Kocan, A.A., Kocan, K.M., Hair, J., 1987, Evidence of a limited schizogonous

cycle for Cytauxzoon felis in bobcats following exposure to infected ticks. J Wildlife Dis

23, 499-501.

77

Bostrom, B., Wolf, C., Greene, C., Peterson, D.S., 2008, Sequence conservation in the rRNA

first internal transcribed spacer region of Babesia gibsoni genotype Asia isolates. Vet

Parasitol 152, 152-157.

Brown, H.M., Berghaus, R.D., Latimer, K.S., Britt, J.O., Rakich, P.M., Peterson, D.S., 2009,

Genetic variability of Cytauxzoon felis from 88 infected domestic cats in Arkansas and

Georgia. J Vet Diagn Invest 21, 59-63.

Brown, H.M., Modaresi, S.M., Cook, J.L., Latimer, K.S., Peterson, D.S., 2009, Genetic

variability of archived Cytauxzoon felis histologic specimens from domestic cats in

Georgia, 1995-2007. J Vet Diagn Invest 21, 493-498.

Brown, H.M., Latimer, K.S., Erikson, L.E., Cashwell, M.E., Britt, J.O., Peterson, D.S., 2008,

Detection of persistent Cytauxzoon felis infection by polymerase chain reaction in three

asymptomatic domestic cats. J Vet Diagn Invest 20, 485-488.

Butt, M.T., Bowman, D., Barr, M.C., Roelke, M.E., 1991, Iatrogenic transmission of Cytauxzoon

felis from a Florida panther (Felix concolor coryi) to a domestic cat. J Wildl Dis 27, 342-

347.

Edwards, A.C., Reichard, M.V., Meinkoth, J.H., Snider, T.A., Meinkoth, K.R., Heinz, R.E.,

Little, S.E., 2010, Confirmation of Amblyomma americanum (Acari: Ixodidae) as a vector

for Cytauxzoon felis (Piroplasmorida: Theileriidae) to domestic cats. J Med Ent.

Ferris, D.H., 1979, A progress report on the status of a new disease of American cats:

cytauxzoonosis. Comp Immunol Microbiol Infect Dis 1, 269-276.

Glenn, B.L., Kocan, A.A., Blouin, E.F., 1983, Cytauxzoonosis in bobcats. J Am Vet Med Assoc

183, 1155-1158.

78

Greene, C.E., Latimer, K., Hopper, E., Shoeffler, G., Lower, K., Cullens, F., 1999,

Administration of diminazene aceturate or imidocarb dipropionate for treatment of

cytauxzoonosis in cats. J Am Vet Med Assoc 215, 497-500, 482.

Haber, M.D., Tucker, M.D., Marr, H.S., Levy, J.K., Burgess, J., Lappin, M.R., Birkenheuer, A.J.,

2007, The detection of Cytauxzoon felis in apparently healthy free-roaming cats in the

USA. Vet Parasitol 146, 316-320.

Harvey, J.W., Dunbar, M.R., Norton, T.M., Yabsley, M.J., 2007, Laboratory findings in acute

Cytauxzoon felis infection in cougars (Puma concolor couguar) in Florida. J Zoo Wildlife

Med 38, 285-291.

Jia, H., Terkawi, M.A., Aboge, G.O., Goo, Y.K., Luo, Y., Li, Y., Yamagishi, J., Nishikawa, Y.,

Igarashi, I., Sugimmoto, C., Fugisaki, K., Xuan, X., 2009, Characterization of a leucine

aminopeptidase of Babesia gibsoni. Parasit 136, 945-52.

Ketz-Riley, C.J., Reichard, M.V., Van Den Bussche, R.A., Hoover, J.P., Meinkoth, J., Kocan,

A.A., 2003, An intraerythrocytic small piroplasm in wild-caught Pallas's cats

(Otocolobus manul) from Mongolia. J Wildlife Dis 39, 424-430.

Kier, A.B., Wagner, J.E., Morehouse, L.G., 1982a, Experimental transmission of Cytauxzoon

felis from bobcats (Lynx rufus) to domestic cats (Felis domesticus). Am J Vet Res 43, 97-

101.

Kier, A.B., Wightman, S.R., Wagner, J.E., 1982b, Interspecies transmission of Cytauxzoon felis.

Am J Vet Res 43, 102-105.

Kocan, A.A., Kocan, K.M., Blouin, E.F., Mukolwe, S.W., 1992, A redescription of schizogony

of Cytauxzoon felis in the domestic cat. Ann Ny Acad Sci 653, 161-167.

79

Kumar, S.K., Tamura, K., Nei, M., 2004, MEGA3: Integrated software for molecular

evolutionary genetics analysis and sequence alignment. Briefings in Bioinformatics 5,

150–163.

Luaces, I., Aguirre, E., Garcia-Montijano, M., Velarde, J., Tesouro, M.A., Sanchez, C., Galka,

M., Fernandez, P., Sainz, A., 2005, First report of an intraerythrocytic small piroplasm in

wild Iberian lynx (Lynx pardinus). J Wildlife Dis 41, 810-815.

Motzel, S.L., Wagner, J.E., 1990, Treatment of experimentally induced cytauxzoonosis in cats

with parvaquone and buparvaquone. Vet Parasitol 35, 131-138.

Nietfeld, J.C., Pollock, C., 2002, Fatal cytauxzoonosis in a free-ranging bobcat (Lynx rufus). J

Wildlife Dis 38, 607-610.

Olivier, C., van de Pas, S., Lepp, P.W., Yoder, K., Relman, D.A., 2001, Sequence variability in

the first internal transcribed spacer region within and among Cyclospora species is

consistent with polyparasitism. Int J Parasitol 31, 1475-1487.

Peixoto, P.V., Soares, C.O., Scofield, A., Santiago, C.D., Franca, T.N., Barros, S.S., 2007, Fatal

cytauxzoonosis in captive-reared lions in Brazil. Vet Parasitol 145, 383-387.

Reichard, M.V., Meinkoth, J.H., Edwards, A.C., Snider, T.A., Kocan, K.M., Blouin, E.F., Little,

S.E., 2008, Transmission of Cytauxzoon felis to a domestic cat by Amblyomma

americanum. Vet Parasitol.

Wagner, J.E., 1976, A fatal cytauxzoonosis-like disease in cats. J Am Vet Med Assoc 168, 585-

588.

Wightman, S.R., Kier, A.B., Wagner, J.E., 1977, Feline cytauxzoonosis: clinical features of a

newly described blood parasite disease. Feline Pract, 24-26.

80

Yabsley, M.J., Murphy, S.M., Cunningham, M.W., 2006, Molecular detection and

characterization of Cytauxzoon felis and a Babesia species in cougars from Florida. J

Wildlife Dis 42, 366-374.

Yamasaki, M., Inokuma, H., Sugimoto, C., Shaw, S.E., Aktas, M., Yabsley, M.J., Yamato, O.,

Maede, Y., 2007, Comparison and phylogenetic analysis of the heat shock protein 70

gene of Babesia parasites from dogs. Vet Parasitol 145, 217-227.

Wenyon, C.M., Hamerton, A.E., 1930, Piroplasms of the West African civet cat (Viverra civetta)

and the Bay lynx (Felis rufa) of North America. Roy Soc Trop Med and Hyg, 24, 7-8.

Xiao, Y.P., Al-Khedery, B., Allred, D.R., 2010, The Babesia bovis VESA1 virulence factor

subunit 1b is encoded by the 1beta branch of the ves multigene family. Mol Biochem

Parasit 171, 81-88.

Zahler, M., Schein, E., Rinder, H., Gothe, R., 1998, Characteristic genotypes discriminate

between Babesia canis isolates of differing vector specificity and pathogenicity to dogs.

Parasitol Res 84, 544-548.

81

CHAPTER 4

NOVEL BABESIA IN A BOBCAT, GEORGIA

Barbara C. Shock, J. Mitchell Lockhart, Adam J. Birkenheuer, and Michael J. Yabsley. 2010. To

be submitted as Letter to the Editor of Emerging Infectious Diseases.

82

To the Editor:

Currently, only two piroplasms have been reported from felines in the North America,

Cytauxzoon felis in domestic cats, bobcats (Lynx rufus), and cougars (Puma concolor) from the

Eastern United States and a novel Babesia species in Florida panthers (P. c. coryii) from

southern Florida (Glenn et al., 1983; Yabsley et al., 2006). During a C. felis surveillance study

involving bobcats (n=799) and cougars (n=49) from thirteen states (Florida, Georgia, Kansas,

Kentucky, Louisiana, Missouri, North Carolina, North Dakota, Ohio, Oklahoma, Pennsylvania,

South Carolina, and West Virginia) a Babesia species was detected in a single female bobcat

from Thomas County, Georgia (n=159; 0.63%).

Sequence analysis of the ITS-1 region indicated that the greatest similarity (92%) was

with a novel large Babesia sp. (Coco) first identified in a domestic dog from North Carolina in

2002 (GenBank accession number: AY618928) (Birkenheuer et al., 2004). The only difference

between the sequence from the bobcat and Babesia sp. (Coco) was that the bobcat Babesia sp.

contained a 45-bp insertion at nucleotide site 434. Based on the identical sequence from bases 1

to 434 and 435 to 557, we believe that this bobcat Babesia sp. represents a variant of Babesia sp.

Coco and not a novel Babesia sp., although additional studies are needed to definitively

determine the con-specificity of these two Babesia. Similar results have been reported with C.

felis where some strains contain a 40 bp insert (Brown et al., 2009a)

To better understand why this bobcat was infected with a variant of Babesia sp. (Coco)

which had only previously been reported from dogs, we conducted additional pathogen screening

using the limited samples available from this trapper-harvested animal. Serum from the bobcat

tested negative for Feline Immunodeficiency virus (FIV) using the IDEXX Snap test. A titer was

1:10 for Feline panleukopenia virus, which was not interpreted as an infection. Interestingly,

83

PCR testing for other parasites revealed that the bobcat was co-infected with C. felis (ITS-2) and

a Bartonella sp. Unfortunately, a blood smear was not available from the Babesia sp. infected

bobcat so that morphologic data could be collected on the Babesia sp. Histological examination

of selected tissues was unrewarding due to the level of necrosis, but Sarcocystis cysts were

observed in muscle tissue. All of these findings are considered incidental and may have

predisposed this bobcat to be more susceptible to Babesia.

Currently, little is known about the natural history of Babesia sp. Coco and the Babesia

sp. detected in the bobcat from Georgia. Babesia sp. Coco was first reported from an

immunosuppressed dog undergoing chemotherapy for lymphoma (Birkenheuer et al., 2004).

Since the initial detection eight additional canine infections have been reported from dogs with

history of travel to Mid-Atlantic states. Six of these dogs were splenectomized and two were

immunosuppressed due to oncolytic drugs (Birkenheuer et al., 2004; Holman et al., 2009;

Sikorski et al., 2010). At least 5 of the 9 dogs infected with Babesia sp. Coco had a history of

tick exposure and at least one sustained bites to the face, a risk factor for other Babesia sp. such

as B. gibsoni (Holman et al., 2009; Yeagley et al., 2009; Sikorski et al., 2010).

Worldwide, several Babesia spp. have been reported from felids including B. herpailuri

and B. pantherae from wild felids in Africa, B. felis and B. leo from domestic cats and wild felids

in Africa, B. cati in domestic cats from India, B. canis canis from domestic cats in Spain, B.

canis presentii from domestic cats in Israel; a Babesia sp. from domestic cats in Portugal,

(Penzhorn et al., 2004; Criado-Fornelio et al., 2004; Jacobson et al., 2000; Baneth et al., 2004).

In the United States, the only report of a Babesia species in a felid is the novel Babesia sp. from

Florida panthers (Yabsley et al., 2006). The panther Babesia sp. appears to be restricted to

Florida panthers because, in the current study, infections were not detected in cougars from TX,

84

LA, GA, or ND. The Babesia sp. from the Florida panther is a small piroplasm and is

indistinguishable from C. felis, whereas Babesia sp. Coco is a large Babesia and based on

sequence analysis of the ITS-1 region, the bobcat Babesia sp. is easily distinguished from the

Florida panther Babesia sp.

In summary, a Babesia sp. closely related to Babesia sp. Coco was detected in a single

bobcat from Georgia which is the first report Babesia infection of a bobcat and the second report

of Babesia in felids from the United States. Based on our data, wild felids, bobcats particularly,

do not likely represent a natural host of Babesia sp. Coco, thus additional surveillance studies are

needed to identify the natural host of this parasite which has been found in immunosuppressed

dogs from numerous southern states.

Acknowledgements

The authors thank numerous personnel from state agencies who collected felid samples.

This study was primarily funded by the Morris Animal Foundation (DO8FE-003) and additional

support was provided by the Federal Aid to Wildlife Restoration Act (50 Stat. 917) and through

sponsorship from fish and wildlife agencies in Alabama, Arkansas, Florida, Georgia, Kansas,

Kentucky, Louisiana, Maryland, Mississippi, Missouri, North Carolina, Oklahoma, Puerto Rico,

South Carolina, Tennessee, Virginia, and West Virginia.

References

Baneth, G., Kenny, M.J., Tasker, S., Anug, Y., Shkap, V., Levy, A., Shaw, S.E., 2004, Infection

with a proposed new subspecies of Babesia canis, Babesia canis subsp. presentii, in

domestic cats. J Clin Microbiol 42, 99-105.

85

Birkenheuer, A.J., Neel, J., Ruslander, D., Levy, M.G., Breitschwerdt, E.B., 2004, Detection and

molecular characterization of a novel large Babesia species in a dog. Vet Parasitol 124,

151-160.

Brown, H.M., Berghaus, R.D., Latimer, K.S., Britt, J.O., Rakich, P.M., Peterson, D.S., 2009,

Genetic variability of Cytauxzoon felis from 88 infected domestic cats in Arkansas and

Georgia. J Vet Diagn Invest 21, 59-63.

Criado-Fornelio, A., Gonzalez-del-Rio, M.A., Buling-Sarana, A., Barba-Carretero, J.C., 2004,

The "expanding universe" of piroplasms. Vet Parasitol 119, 337-345.

Glenn, B.L., Kocan, A.A., Blouin, E.F., 1983, Cytauxzoonosis in bobcats. J Am Vet Med Assoc

183, 1155-1158.

Holman, P.J., Backlund, B.B., Wilcox, A.L., Stone, R., Stricklin, A.L., Bardin, K.E., 2009,

Detection of a large unnamed Babesia piroplasm originally identified in dogs in North

Carolina in a dog with no history of travel to that state. J Am Vet Med Assoc 235, 851-

854.

Jacobson, L.S., Schoeman, T., Lobetti, R.G., 2000, A survey of feline babesiosis in South Africa.

J S Afr Vet Assoc 71, 222-228.

Penzhorn, B.L., Schoeman, T., Jacobson, L.S., 2004, Feline babesiosis in South Africa: a review.

Ann N Y Acad Sci 1026, 183-186.

Sikorski, L.E., Birkenheuer, A.J., Holowaychuk, M.K., McCleary-Wheeler, A.L., Davis, J.M.,

Littman, M.P., 2010, Babesiosis caused by a large Babesia species in 7

immunocompromised dogs. J Vet Intern Med 24, 127-131.

86

Yabsley, M.J., Murphy, S.M., Cunningham, M.W., 2006, Molecular detection and

characterization of Cytauxzoon felis and a Babesia species in cougars from Florida. J

Wildlife Dis 42, 366-374.

Yeagley, T.J., Reichard, M.V., Hempstead, J.E., Allen, K.E., Parsons, L.M., White, M.A., Little,

S.E., Meinkoth, J.H., 2009, Detection of Babesia gibsoni and the canine small Babesia

'Spanish isolate' in blood samples obtained from dogs confiscated from dogfighting

operations. J Am Vet Med Assoc 235, 535-539.

87

CHAPTER 5

CONCLUSIONS

Cytauxzoon felis, a protozoan parasite of wild and domestic felids, is the causative agent

of cytauxzoonosis in domestic and some exotic felids in the United States. Previous research has

identified the bobcat (Lynx rufus) as the natural reservoir for this parasite. C. felis has been

experimentally transmitted by two tick species, Dermacentor variabilis and Amblyomma

americanum, which have overlapping distributions throughout the southeastern United States.

Part of the current study was to determine the distribution and prevalence of C. felis in

free-ranging bobcat populations from 13 states including California, Colorado, Florida, Georgia,

Kansas, Kentucky, Missouri, North Carolina, North Dakota, Ohio, Oklahoma, South Carolina,

and West Virginia. These states were selected because D. variabilis is present in each state, but

A. americanum is only currently known to be present in a subset of these states. Blood or spleen

samples from 697 trapper-harvested or road-killed bobcats were tested for C. felis infection by a

polymerase chain reaction (PCR) assay which targets the first ribosomal internal transcribed

spacer region (ITS-1). Significantly higher C. felis prevalences were detected from Missouri

(79%, n=39), North Carolina (63%, n=8), Oklahoma (60%, n=20), South Carolina (57%, n=7),

Kentucky (55%, n=74), Florida (44%, n=45), and Kansas (27%, n=41) compared with Georgia

(9%, n=159), North Dakota (2.4%, n=124), Ohio (0%, n=19), and West Virginia (0%, n=37). In

addition to bobcats, we tested a limited number of samples (n=8) from other wild felids and only

one cougar (Puma concolor) from Louisiana was positive, which represents the first report in an

88

infected cougar outside the Florida panther population. These data also indicate that C. felis is

present in North Dakota where C. felis has not been reported in domestic cats. Based on linear

regression analysis, prevalence rates were significantly higher in states where there are

established populations of A. americanum, which supports recent data on the experimental

transmission of C. felis by A. americanum and the fact that domestic cat cases are temporally

associated with A. americanum activity. Collectively, these data confirm that bobcats are a

common reservoir for C. felis and that A. americanum likely is an epidemiologically important

vector.

Interestingly, we detected a single bobcat from Georgia that was infected with a Babesia

sp. which is the first report Babesia infection of a bobcat and the second report of a Babesia sp,

in felids from the United States. The Babesia sp. was closely related to Babesia sp. Coco which

has only been reported from 9 immunosuppressed domestic dogs. Based on our data, wild felids,

bobcats particularly, do not likely represent a natural host of Babesia sp. Coco, thus additional

surveillance studies are needed to identify the natural host of this parasite.

The next goal of this project was to genetically characterize select samples of C. felis

circulating in wild felids. Although most domestic cats infected with the parasite die of clinical

illness, increasing numbers of cats have been identified surviving infection. Because the bobcat

is the natural reservoir, they typically only develop limited or no clinical signs of illness.

Previous studies, including our own, show that in many areas, free-ranging bobcats have high

prevalence rates. Although not likely important reservoirs, C. felis has also been detected in free-

ranging Florida panthers. Recent studies have identified specific genotypes of C. felis circulating

in domestic cats which are associated with different clinical outcomes and specific spatial

locations. In the current study, we investigated the intraspecific variation of the C. felis internal

89

transcribed spacer (ITS)-1 and ITS-2 rRNA regions from 153 bobcats and eight cougars from 11

states (Florida, Georgia, Kansas, Kentucky, Louisiana, Missouri, North Carolina, North Dakota,

South Carolina, Oklahoma, and Pennsylvania). Unambiguous ITS-1 and ITS-2 data were

obtained for 131 and 106 samples, respectively, and both ITS-1 and ITS-2 sequences were

obtained for 82 (51%) samples. Within the ITS-1 region, sequences from 46 bobcats were

identified which were identical to those previously reported in domestic cats and we identified 60

unique sequences. Samples from 46 bobcats and one cougar had ITS-1 sequences identical to the

most common sequence reported from domestic cats. The most common ITS-2 sequence from

domestic cats was also common in wild felids (36 bobcats and a cougar). Five other ITS-2

sequences had been previously reported in domestic cats; the remaining ITS-1 and ITS-2

sequences from wild felids were novel. Samples from three Florida panthers and a bobcat from

MO had a 40/41-bp insert in the ITS-2 similar to one described previously in a cat from AR.

Additionally, a previously undescribed 198/199-bp insert was detected in the ITS-2 sequence

from in four bobcats. Collectively, nine bobcats and one cougar were infected with genotype

ITSA and two bobcats were infected with genotypes ITSB and ITSC. Eight bobcats from

southwestern Georgia and northwestern Florida were infected with a novel genotype (ITSD).

These data indicate that based on ITS1 and ITS2 sequences, numerous C. felis strains circulate in

wild felids and that some genotypes may cluster in specific spatial locations.

Collectively, these studies indicate that the prevalence of C. felis is high in many

populations of bobcat, especially in those where A. americanum densities are high compared

with D. variabilis. Genetic characterization data indicate that C. felis samples from wild felids

has significantly more variability in the ITS-1 and ITS-2 regions compared with domestic cats.

This is probably attributable to the long relationship between this parasite and its natural wild

90

mammalian hosts, as domestic cats are spill-over or accidental hosts for the parasite. These

studies also revealed the infection of a bobcat with a novel species of Babesia only previously

reported from immunocompromised dogs, which is the first report of a Babesia sp. in bobcats

and the second report of a Babesia sp. in a felid from the United States. The current study is the

most comprehensive study on piroplasms in free-ranging wild felid populations in the United

States.

Future research into C. felis in bobcat populations should focus on areas where A.

americanum is expanding into naïve populations of wild felids. A majority of the bobcats in this

study were presumably healthy free-ranging felids, but L. rufus has been known to die of

cytauxzoonosis. C. felis could be a selection force on bobcat populations in areas such as the

American West and states such as Ohio and West Virginia, where C. felis has not been reported

in domestic or wild felids. Population level studies could address the dynamics of infection and

survival as C. felis emerges in these areas.

Another way to understand the role of C. felis in bobcat populations would be to address

the bobcat population itself. A genetics study on bobcats could reveal if there are subspecies or

types of bobcats which are more susceptible to infection with C. felis. In addition, a long-term

tracking study could investigate the immune responses in felids infected and uninfected with C.

felis. Although C. felis causes little to no harm to bobcat populations currently, with the rise of

feral cat populations in the United States, these wild felids have increasing contact with new

diseases. It will be important to understand the role of C. felis in bobcat populations as their

exposure to novel health threats increases.