MOLECULAR AND BIOLOGIC CHARACTERIZATION OF TRYPANOSOMA CRUZI
FROM THE UNITED STATES OF AMERICA
by
DAWN MARIE ROELLIG
(Under the Direction of Michael J. Yabsley)
ABSTRACT
Trypanosoma cruzi, the causative agent of Chagas disease, is a flagellated protozoan parasite endemic to the Americas. There are an estimated 7.6 million people infected within Latin America, with 200,000 new cases occurring annually. In the United
States, only six autochthonous human cases have been reported, but screening of US blood donors since January 2007 by the Chagas’ Disease Biovigilance Network has confirmed over 1,000 seropositive donations, some of which are believed to be autochthonously acquired infections. In addition to autochthonous human cases, naturally acquired infections have been reported in domestic dogs, captive exotic animals, and a wide range of wildlife species. Although T. cruzi is common in the sylvatic cycle
(transmission between wildlife and vectors) and the peridomestic cycle (between animals that move between the wild and around homes), circulation within the domestic cycle
(between hosts within or around homes) is less often documented in the US. In the sylvatic cycle, a number of wild mammals have been identified as reservoirs and the prevalence of T. cruzi in US wildlife based on serology, culture isolation, and/or PCR can be equally as high as in South America. Considerable work on T. cruzi has been
conducted in Central and South America, but due to limited numbers of human cases in the US, little work has been conducted to characterize US isolates. Previous studies have determined prevalence in some host species through the use of serology, hemoculture, and/or PCR, identified transmission routes for a few reservoirs using non-native isolates, and molecularly characterized a limited number of isolates. With new human cases, increasing numbers of veterinary cases, and influx of potentially infected immigrants, understanding the ecology in the US is imperative. The goal of this dissertation was to elucidate characteristics of T. cruzi isolates from the US using molecular and in vivo techniques to better understand T. cruzi epizootiology in the region.
INDEX WORDS: Trypanosoma cruzi, trypanosome, Chagas disease, vector-borne disease, zoonotic disease, experimental infection, United States, sylvatic cycle
MOLECULAR AND BIOLOGIC CHARACTERIZATION OF TRYPANOSOMA CRUZI
FROM THE UNITED STATES OF AMERICA
by
DAWN MARIE ROELLIG
BA, Agnes Scott College, 2004
MS, Georgia Southern University, 2006
A Dissertation Submitted to the Graduate Faculty of The University of Georgia in Partial
Fulfillment of the Requirements for the Degree
DOCTOR OF PHILOSOPHY
ATHENS, GEORGIA
2009
© 2009
Dawn Marie Roellig
All Rights Reserved
MOLECULAR AND BIOLOGIC CHARACTERIZATION OF TRYPANOSOMA CRUZI
FROM THE UNITED STATES OF AMERICA
by
DAWN MARIE ROELLIG
Major Professor: Michael J. Yabsley
Committee: David S. Peterson Ray M. Kaplan David E. Stallknecht Daniel G. Mead
Electronic Version Approved:
Maureen Grasso Dean of the Graduate School The University of Georgia December 2009
iv
ACKNOWLEDGEMENTS
I would like to extend a special thank you to Dr. Michael Yabsley for all his support, guidance, and patience through this project and my academic career. My entire committee, including Dr. David Peterson, Dr. Ray Kaplan, Dr. David Stallknect, Dr.
Daniel Mead, and Dr. Michael Yabsley, has given me invaluable advice, encouragement, education, and critique for which I am grateful. I would also like to thank my mentors throughout my undergraduate and graduate education, Dr. Robert Massung, Dr. Dana
Nayduch, and Dr. Lance Durden, all of whom were instrumental in introducing me to research, parasitology, and infectious diseases.
I offer a sincere thanks to my parents, Ralph and Darlene Roellig, who instilled in me the courage to never give up, sound advice, and ice cream when I needed it most.
Many thanks to all the faculty, staff, and students at the Southeastern Cooperative
Wildlife Disease Study for all their assistance and support throughout the years. I especially would like to thank all those who have previously worked in the Yabsley lab that have helped with the various experiments and projects, including Letitia Saunders,
Mason Savage, Wendy Fujita, Kate McMillan, Emily Rushton, Jessica Murdock, Barbara
Shock, Emily Blizzard, Jess Gonynor, and Elizabeth Gleim. A gigantic thank you to all the friends I’ve made over the years in Athens, especially Lindsay Fann Killmaster and
Sabrina McGraw who have been there for me through it all.
TABLE OF CONTENTS
Page
ACKNOWLEDGEMENTS ...... iv
LIST OF TABLES ...... ix
LIST OF FIGURES ...... xii
CHAPTER
1 INTRODUCTION ...... 1
References ...... 5
2 LITERATURE REVIEW ...... 11
History, life cycle and epidemiology of Trypanosoma cruzi ...... 11
Human cases of Trypanosoma cruzi in the United States...... 15
Trypanosoma cruzi vectors in the United States...... 17
Trypanosoma cruzi in wildlife from the United States ...... 17
Trypanosoma cruzi diagnostics...... 20
Trypanosoma cruzi molecular evolution and genotyping ...... 20
Wildlife reservoirs and the genotype-strain dichotomy ...... 24
References ...... 28
3 MOLECULAR TYPING OF TRYPANOSOMA CRUZI ISOLATES, UNITED
STATES ...... 43
Abstract ...... 44
The Study ...... 45
vi
Conclusion ...... 46
Acknowledgments...... 48
Biographical Sketch ...... 48
References ...... 49
4 GENETIC VARIATION AND EXCHANGE IN TRYPANOSOMA CRUZI
ISOLATES FROM THE UNITED STATES ...... 55
Introduction ...... 57
Materials and Methods ...... 59
Results ...... 61
Discussion ...... 64
Acknowledgements ...... 68
References ...... 68
5 GENETICALLY DIFFERENT ISOLATES OF TRYPANOSOMA CRUZI
ELICIT DIFFERENT INFECTION DYNAMICS IN RACCOONS
(PROCYON LOTOR) AND VIRGINIA OPOSSUMS (DIDELPHIS
VIRGINIANA) ...... 86
Abstract ...... 87
Introduction ...... 88
Materials and Methods ...... 90
Results ...... 95
Discussion ...... 99
Acknowledgements ...... 104
References ...... 105
vii
6 EXPERIMENTAL INFECTIONS OF TWO SOUTH AMERICAN
RESERVOIRS WITH FOUR DISTINCT STRAINS OF TRYPANOSOMA
CRUZI ...... 119
Summary ...... 120
Introduction ...... 121
Materials and Methods ...... 123
Results ...... 126
Discussion ...... 129
Acknowledgements ...... 132
Financial Support ...... 133
Literature Cited ...... 133
7 ORAL TRANSMISSION OF TRYPANOSOMA CRUZI WITH OPPOSING
EVIDENCE FOR THE THEORY OF CARNIVORY ...... 143
Abstract ...... 144
Materials and Methods ...... 146
Results ...... 150
Discussion ...... 151
Acknowledgments...... 154
Literature Cited ...... 154
8 EVALUATION OF THE CHAGAS STAT-PAK ™ ASSAY FOR
DETECTION OF TRYPANOSOMA CRUZI ANTIBODIES IN WILDLIFE
RESERVOIRS ...... 161
Abstract ...... 162
viii
Literature Cited ...... 168
9 INFECTIVITY OF SYLVATIC AND DOMESTIC DOG TRYPANOSOMA
CRUZI ISOLATES FROM THE UNITED STATES TO LABORATORY
MICE AND RATS ...... 175
Abstract ...... 176
Acknowledgements ...... 182
Financial Support ...... 182
References ...... 183
10 CONCLUSIONS...... 189
Study 1 (Chapter 3) ...... 189
Study 2 (Chapter 4) ...... 190
Study 3 (Chapter 5) ...... 191
Study 4 (Chapter 6) ...... 192
Study 5 (Chapter 7) ...... 192
Study 6 (Chapter 8) ...... 193
Study 7 (Chapter 9) ...... 194
References ...... 194
ix
LIST OF TABLES
Page
Table 2.1: Wildlife species fund to be naturally infected with Trypanosoma cruzi through culture or direct observation of the parasite or serologic detection of antibodies reactive to T. cruzi ...... 26
Table 3.1: Origin and lineage identification of the 107 United States isolates used in the study ...... 52
Table 3.2: Approximate amplicon sizes of gene targets and lineage determination ...... 54
Table 4.1: Lineage typing of Trypanosoma cruzi isolates from the United States ...... 76
Table 4.2: Nucleotide sequence variations within the MSH2 gene sequence of 50 T. cruzi isolates from the United States compared to reference strains ...... 78
Table 4.3: Nucleotide sequence variation within the Tc52 gene sequence of 51 T. cruzi isolates from the United States compared to reference strains ...... 79
Table 5.1: Results of polymerase chain reaction (PCR) amplification of Trypanosoma cruzi 24SαrDNA D7 divergent domain indirect immunofluorescence assay (IFA), and hemoculture from experimentally infected raccoons ...... 111
Table 5.2: Polymerase chain reaction (PCR) amplification of Trypanosoma cruzi minicircle gene in tissues collected at necropsy from experimentally infected raccoons112
Table 5.3: Inflammation scores of tissues collected at necropsy from experimentally infected raccoons ...... 113
x
Table 5.4: Results of polymerase chain reaction amplification (PCR) of Trypanosoma cruzi minicircle, indirect immunofluorescence assay (IFA), and hemoculture from experimentally infected Virginia opossums...... 114
Table 5.5: Polymerase chain reaction (PCR) amplification of Trypanosoma cruzi minicircle gene in tissues collected at necropsy from experimentally infected Virginia opossums ...... 115
Table 5.6: Inflammation scores of tissues collected at necropsy from experimentally infected Virginia opossums...... 116
Table 6.1: Results of polymerase chain reaction (PCR) amplification of Trypanosoma cruzi 24Sα rDNA D7 divergent domain and indirect immunofluorescence assay (IFA) from experimentally infected degus...... 139
Table 6.2: Results of polymerase chain reaction amplification (PCR) of Trypanosoma cruzi 24Sα rDNA D7 divergent domain and indirect immunofluorescence assay (IFA) of experimentally infected short-tailed opossums ...... 140
Table 7.1: Inflammatory lesions of tissues in raccoons experimentally infected with
Trypanosoma cruzi via different routes ...... 159
Table 8.1: Results of hemoculture, indirect immunofluorescent antibody (IFA), and
Chagas Stat-Pak™ assay testing of 57 wild raccoons from Georgia and Florida ...... 173
Table 8.2: Indirect immunofluorescent antibody (IFA) and Chagas Stat-Pak™ testing results for raccoons (Procyon lotor) and degus (Octodon degu) experimentally-infected with Trypanosoma cruzi ...... 174
Table 9.1: Detection of Trypanosoma cruzi in eight acutely-infected and one chronically- infected Crl:CD1 (ICR) mice ...... 187
xi
Table 9.2: Detection of Trypanosoma cruzi in three acutely-infected and one chronically- infected white laboratory mice ...... 188
xii
LIST OF FIGURES
Page
Figure 2.1: Life cycle of Trypanosoma cruzi...... 14
Figure 2.2: Distribution of confirmed seropositive donations to the US blood bank as reported by the Chagas’ Biovigilance network ...... 16
Figure 2.3: Classic typing scheme of Trypanosoma cruzi ...... 22
Figure 4.1: Evolutionary relationships among e mismatch-repair class 2 gene (MSH2) and the thiol-disulfide oxido-reductase Tc52 gene (Tc52) from 50 and 51 Trypanosoma cruzi isolates, respectively ...... 80
Figure 4.2: Evolutionary relationships among dihydrofolate reductase-thymidylate synthase (DHFR-TS) from 43 Trypanosoma cruzi isolates ...... 82
Figure 4.3: Evolutionary relationships among cytochrome oxidase subunit II- NADH dehydrogenase subunit I region (COII-ND1) from 43 Trypanosoma cruzi isolates ...... 84
Figure 5.1: Parasitemias of individual raccoons (Procyon lotor) experimentally inoculated with different genotypes of Trypanosoma cruzi ...... 117
Figure 5.2: Parasitemias of individual Virginia opossums (Didelphis virginiana) experimentally inoculated with TcI and dual (IIa & I) genotypes of Trypanosoma cruzi118
Figure 6.1: Parasitemias of degus (Octodon degus) experimentally inoculated with different genotypes of Trypanosoma cruzi ...... 141
Figure 6.2: Parasitemias of short-tailed opossums (Monodelphis domestica) experimentally inoculated with different genotypes of Trypanosoma cruzi ...... 142
xiii
Figure 7.1: Parasitemias of raccoons experimentally infected with Trypanosoma cruzi via different inoculation methods ...... 160
1
CHAPTER 1
INTRODUCTION
The ultimate goal of this dissertation was to identify molecular and biological characteristics of Trypanosoma cruzi in the United States (US) and associate such characters with infection dynamics and virulence in mammalian hosts. T. cruzi is a flagellated protozoan parasite endemic to the Americas. An estimated 7.6 million people within Latin America are infected with the parasite and 200,000 new cases occur annually (CDC, 2010; TDR, 2005). Transmission occurs with the invasion of the infective metacyclic stage, which is passed in the feces of blood-feeding, triatomine vectors, at a mucous membrane or cut or abrasion at the bug bite wound. Once infected, an individual can develop Chagas‘ disease (Chagas, 1909), also known as American
Trypanosomiasis. Manifestations of infection vary, but acute and chronic stages of disease occur. During the acute phase, individuals may experience fever, edema, general malaise, and a characteristic chagoma at the site of parasite entry. Chronic Chagas‘ disease is a result of tissue damage caused by the intracellular form of T. cruzi and can include cardiomyopathy, megacolon, and/or megaesophus.
In the United States, only six autochthonous human cases have been reported
(Woody and Woody, 1955; Schiffler et al, 1984; Ochs et al, 1996; Herwaldt et al, 2001;
Dorn et al, 2007), but screening of US blood donors from January 2007 to November
2009 by the Chagas‘ Disease Biovigilance Network has confirmed 1,062 seropositive 2 donations (AABB, 2009), some of which are believed to be autochthonously acquired infections. In addition to autochthonous human cases, naturally acquired infections have been reported in domestic dogs, captive exotic animals, and a wide range of wildlife species in the United States (Kjos et al., 2008; Williams et al., 2009, Brown et al., in press). Although circulation within the domestic cycle (between hosts within or around homes) is less often documented in the US, T. cruzi is common in the sylvatic cycle
(transmission between wildlife and vectors) and the peridomestic cycle (between animals that move between the wild and around homes). In the sylvatic cycle, a number of wild mammals have been identified as reservoirs and the prevalence of T. cruzi in US wildlife based on serology, culture isolation, and/or PCR can be equally as high as in South
America (Barr et al, 1991).
Genotyping of T. cruzi has led to the acceptance of six phylogenetic lineages (TcI,
TcIIa thru TcIIe) as molecular characters of the parasite (Brisse et al., 2000; Brisse et al.,
2001). While all six genotypes have been detected in mammals from South America, previous work indicated two T. cruzi genotypes (TcI and TcIIa) were present in the US
(Clark and Pung, 1994; Briones et al, 1999). Previous investigations have also suggested a host-genotype association (Clark and Pung, 1994; Briones et al, 1999; Yeo et al, 2005), but these studies were limited in relevance to US T. cruzi strains due to low sample numbers, low host diversity, and narrow geographic distribution (Miles et al, 1978; Clark and Pung, 1994; Barnabé et al, 2001; Yabsley and Noblet, 2002; Yeo et al, 2005). In particular, it has been hypothesized that marsupials are better hosts for TcI strains while placental mammals are better hosts for TcII strains (Yeo et al., 2005). Experimental investigations testing the susceptibility of these two different mammal infraclasses to 3 different T. cruzi genotypes have not been undertaken and evidence is only presented in surveillance studies (Clark and Pung, 1994; Briones et al, 1999; Yeo et al, 2005; Roque et al., 2008; Rozas et al.. 2007; Spotorno et al., 2008; Galuppo et al., 2009).
In addition to molecular characterization by genotyping, strains of T. cruzi have been used to illustrate rare genetic exchange events in the normally clonal population
(Machado and Ayala, 2001; Brisse et al., 2003; Gaunt et al., 2003). However, these studies were conducted with South American isolates, and few have investigated genetic diversity and recombination in North American isolates (Clark and Pung, 1994; Barnabé et al, 2001; Machado and Ayala, 2001; Iwagami et al., 2007; Subileau et al., 2009).
Studies of US isolates in a similar manner may reveal clues for the evolutionary ecology of T. cruzi in the region; it is suggested that geographical isolation of North and Central
America from South America allowed isolation of placentals and marsupials and independent evolution of T. cruzi infecting either group of mammals (Briones et al.,
1999).
Low T. cruzi infection rates of humans in the United States are likely indicative of different living conditions compared to regions where the parasite is endemic. For example, the vector feeds at night and dwells in cracks and holes in homes during the day; many houses in South America are poorly constructed, providing a habitat for the triatomes. However, the virulence and infectivity of T. cruzi in the United States may differ greatly from South American isolates and cannot be ruled out as a factor in the low prevalence. Heterogeneity of two nuclear gene targets, Tc52 and MSH2, has been suggested to alter virulence between T. cruzi strains from South America previously
(Ouaissi et al., 1995a; Ouaissi et al., 1995b; Oury et al., 2005; Mathieu-Daudé et al., 4
2007; Augusto-Pinto et al., 2001). Investigations of heterogeneity between US T. cruzi isolates are limited (Clark and Pung, 1994) and a difference in infectivity between genotypes has not been investigated.
T. cruzi is a genetically and biologically diverse parasite, as is evident with work conducted with South American isolates. Although the presence of T. cruzi in the US is known, biological and molecular characteristics in the region are understudied. Taking into account the limited data previously presented for the United States, the goal of this dissertation was to identify molecular and biological characteristics of T. cruzi in the region. The central hypothesis to this research is that distinct genotypes (TcI or TcIIa) of
T. cruzi have different hosts and transmission cycles and have different biological characteristics. To test this hypothesis a series of specific aims are presented
1. Genetically classify autochthonously acquired US T. cruzi from free-ranging and
captive wildlife, domestic animals, triatomine bug vectors, and humans.
2. Determine the infection dynamics and transmission alternatives for natural
wildlife reservoirs from the US and South America experimentally infected with
T. cruzi isolates from US wildlife.
3. Based on molecular characters, evaluate efficacy differences in rapid T. cruzi
testing of samples from US and South American T. cruzi wildlife reservoirs.
4. Determine pathogenicity and virulence of genetically classified isolates of T. cruzi
from the US in murine models.
5
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Augusto-Pinto, L., Bartholomeu, D.C., Teixeira, S.M.R., Pena, S.D.J., Machado, C.R.
2001. Molecular cloning and characterization of the DNA mismatch repair gene
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Barr SC, Brown CC, Dennis VA, Klei TR. 1991. The lesions and prevalence of
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Briones MRS, Souto RP, Stolf BS, Zingales B.1999. The evolution of two Trypanosoma
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Brisse D, Barnabé C, Tibayrenc M. 2000. Identification of six Trypanosoma cruzi
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MRC, Buck GA, Dujardin JC, Tibayrenc M. 2003. Evidence for genetic exchange
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Brown, EL, Roellig, DM, Gompper, ME, Monello, RJ, Wenning, KM, Gabriel, MW,
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Iwagami M, Higo H, Miura S, Yanagi T, Tada I, Kano S, Agatsuma T. 2007. Molecular
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Coexistence of Trypanosoma cruzi genotypes in wild and peridomestic mammals
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11
CHAPTER 2
LITERATURE REVIEW
History, life cycle and epidemiology of Trypanosoma cruzi
Trypanosoma cruzi is the causative agent of Chagas‘ disease or American
Trypanosomiasis. It is a hemoflagellated, protozoan parasite within the class
Kinetoplastida. The characteristic morphological character of this group is a kinetoplast, a dark-staining structure composed of extranuclear DNA that is present within the parasite‘s mitochondrion. T. cruzi was first discovered in the midgut of Triatoma infestans, a reduviid bug, by Carlos Chagas while he served as physician at a railroad stop in Lassance, Brazil (Bastien, 1998). Soon after, he examined an ill cat and found the presence of T. cruzi in its blood; weeks later at the same home, Chagas examined a febrile three-year-old child that had been bitten by reduviids. The young girl exhibited trypanosomes in her peripheral blood, splenomegaly, myxedema, and eventually died of the acute phase of Chagas‘ disease. Within the year, Carlos Chagas had discovered the parasite, vector, and hosts (Chagas, 1909).
The basic life cycle of T. cruzi was first described upon its discovery by Chagas in
1909 (Figure 2.1). T. cruzi utilizes stercorarian transmission, unlike many other pathogenic trypanosomes (e.g., T. brucei) that utilize salivarian transmission. Within an infected vector, epimastigotes proliferate in the reduviid midgut then migrate towards the hindgut and attach to the wall hydrophobically prior to differentiation into metacyclic 12 trypomastigotes (Bonaldo et al., 1988; Kleffman et al., 1998). The metacyclic forms are excreted with the bug‘s feces and are infective to the mammalian host. The parasite enters the bloodstream by invading at a mucous membrane or cut or abrasion at the bite wound. Within the mammalian host, metacyclics can invade any nucleated cell and, once inside a cell, differentiate to amastigotes. This intracellular form replicates by binary fission and differentiates to blood stream forms that exit a cell once the cytoplasm is full of parasites. This infective stage will circulate in the blood until ingested by a reduviid bug during a blood meal. Of the various morphological forms of T. cruzi, the amastigotes, sphaeromastigotes (an intermediate form between amastigotes and epimastigotes or epimastigotes and trypomastigotes) and epimastigotes are replicative (Tyler and Engman,
2001).
Trypanosoma cruzi is endemic to the Americas, particularly Central and South
America, with approximately 7.6 million people infected (CDC, 2010). Nearly 200,000 new cases occur annually, with transmission via classical transmission by a vector, oral transmission by ingestion of vector parts in contaminated food or drink, or vertical transmission from mother to offspring (Hoff et al., 1978; TDR, 2005; Ianni and Mady,
2005; Muños et al., 2007). Cases may occur in non-endemic regions (e.g. United States,
Canada, Europe, Japan, and Australia) as a result of international travel to endemic regions, immigration of infected individuals, or infection acquired through organ transplant or blood transfusion (Schmunis, 2007; Steele et al., 2007; Lescure et al., 2008;
Gascon et al., 2009; Bern and Montgomery, 2009).
Infected individuals may exhibit clinical signs characteristic of the acute and chronic stage of disease, only one stage, or neither stage. Fever, edema, general malaise, 13 hepatosplenomegaly, muscle pains, sweats, anorexia, irritability and sometimes vomiting and diarrhea may present during the acute stage (Santos-Buch and Acosta, 1985). Also a characteristic chagoma will form at the site of parasite entry; if the entry site is the conjunctiva, edema around the eye, called Romaña‘s sign, is observed. Higher incidence of acute Chagas‘ disease occurs in children under the age of five (Santos-Buch and
Acosta, 1985). The acute stage of disease can last up to 90 days and is followed by an asymptomatic period before signs of chronic Chagas are exhibited (CDC, 2010).
Approximately 70-80% of individuals remain in this indeterminate stage for the remainder of their lives, and 20-30% of infected individuals will develop chronic Chagas‘ disease (Pinto Dias, 1995; Maguire et al., 1987). Commonly, during chronic Chagas‘ disease, the intracellular form of the parasite, the amastigote, causes damage to the tissue in which it invades. Within cells that typically show the greatest signs of pathology, such as smooth, skeletal, and cardiac muscle, changes in expression levels of important trafficking proteins occur. As a result, there is diminished uptake of normal nutrients and endocytic ligands, which prevents the parasites from being cleared from the cell (Batista et al., 2006). This occurs because the cells are weakened by a lack of proper nutrients and function is impeded by parasites residing within the cell. Additionally in cardiac muscle, cytoskeleton elements remodel within an infected cell (Barbosa and Meirelles, 1995) and myofibrils breakdown and rearrange in a loose network (Pereira et al., 1993). T. cruzi- infected cells will weaken due to these changes, and impaired contractility has been noted in vivo because of them (Taniwaki et al., 2006). These cellular changes during the chronic stage can result in megacolon, megaesophagous, and cardiomyopathy. 14
Figure 2.1. Life cycle of Trypanosoma cruzi 15
Human cases of Trypanosoma cruzi in the United States
Prior to detection of autochthonous human cases in the United States, the pathogenicity of a Texas isolates of T. cruzi was demonstrated in an experimental human infection that resulted in clinical signs identical to that of South American cases
(Packchanian, 1943). The first autochthonous human case was diagnosed in a 10-month- old child from Corpus Christi, TX displaying a fever, irritability, and swollen eyelids
(Woody and Woody, 1955). Initially thought to be an unknown viral infection or leukemia, a blood smear from the child revealed trypomastigotes and a diagnosis of T. cruzi infection was confirmed (Woody and Woody, 1955). Since the first case, five others have been reported from California, Texas, Tennessee, and Louisiana (Schiffler et al.,
1984; Ochs et al., 1996; Herwaldt et al., 2001; Dorn et al., 2007).
Previous serologic surveys within the United States have been conducted in
Arizona, California, Georgia, and Texas. On Indian reservations in Arizona, antibody prevalence ranged from 1% in the Pima Indians (J Miller, personal communication) to
4.4% in Papago Indians (Miller et al., 1977). A survey of the area surrounding the 1982
California human case revealed six of 237 individuals tested were seropositive for T. cruzi (Navin et al., 1985). A Georgia study tested 951 patients and detected antibodies in four individuals (Farrar et al., 1963). Testing of 122 myocardial disease patients in
Georgia detected two weakly seropositive individuals (Farrar et al., 1972). A follow-up serosurvey surrounding the first two human cases in southern Texas showed prevalence of 1.4% in children (Woody et al., 1965). Additionally, 2.5% of 117 individuals bitten by
T. gerstaeckeri in Texas were seropositive (Woody et al., 1965). 16
Since January 2007, serologic testing of the American Red Cross Blood Bank has revealed over 1,000 T. cruzi seropositive submissions (AABB, 2009). The distribution of seropositive donations is nationwide, but most come from Florida, Texas, and California, all of which are states in which human autochthonous cases have previously been reported (Figure 2.2). It has been estimated that 300,167 individuals are infected with T. cruzi in the United States based on antibody prevalence data for immigrant populations
(Bern and Montgomery, 2009). Both studies highlight the need for data collection that will reveal the impact T. cruzi has in the United States.
Figure 2.2. Distribution of confirmed seropositive donations to the US blood bank as reported by the Chagas’ Biovigilance network (AABB, 2009). 17
Trypanosoma cruzi vectors in the United States
T. cruzi in the United States was first detected in a vector, Triatoma protracta, collected from wood rat (Neotoma spp.) nests in San Diego, California (Kofoid and
McCulloch, 1916). Additional infected vector species within the United States included
T. sanguisuga, T. lecticularia, T. gerstaeckeri, T. neotomae, T. recurva and T. rubida
(Ryckman et al., 1965). Of these potential vectors, T. sanguisuga has the widest distribution across the southeast ranging from Maryland to Texas (Lent et al., 1979).
However, the accurate distribution of many of these vector species is poorly defined
(Kjos et al., 2009). T. cruzi-infected triatomes have been observed in Georgia (Pung et al., 1995), Florida (Beard et al., 1998), Alabama (Olsten et al., 1964), Tennessee
(Herwaldt et al., 2000), and Louisiana (Yaeger, 1961). High prevalences of T. cruzi infection in vectors have been reported in Texas, California, and Arizona at 17-50.74%,
14-40%, 7.1-20.5%, respectively (Burkholder et al. 1980; Kjos et al., 2009; Pippin 1970;
Ryckman and Ryckman 1967; Sullivan et al. 1949; Wood 1949, 1975; Wood and Wood
1964). The implications for human infections is unknown (Kjos et al., 2009), but it is known that T. protracta was implicated in the Tennessee human case, and T. sanguisuga heavily infested the home and surrounding property of the 2006 Louisiana case (Herwaldt et al., 2000; Dorn et al., 2007).
Trypanosoma cruzi in wildlife from the United States
In comparison to both vector and human prevalence, wildlife within the United
States appears to have much higher prevalence rates (Table 2.1). The first reports of natural T. cruzi infections in the United States occurred in the early 1940s from wildlife 18
(nine-banded armadillo (Dasypus novemcinctus), Virginia opossum (Didelphis virginiana), pallid bat (Antrozous pallidus), house mouse (Mus musculus), and three species of wood rats (Neotoma spp.)) (Usinger, 1944). Raccoons were later identified as reservoirs (Walton et al, 1956). During a study of leptospirosis surveillance, T. cruzi-like organisms were isolated from the urine and/or kidney of gray foxes (Urocyon cinereoargenteus) and striped skunks (Mephitis mephitis) (McKeever et al., 1958).
Prevalence based on serology and/or culture in raccoons has ranged from 1.5% in southwestern Georgia and northwestern Florida (McKeever et al, 1958) to 63% in
Oklahoma (John and Hoppe, 1986). In Virginia opossums, prevalence ranges from 8% in
North Carolina (Karsten et al., 1992) to 33% in southern Louisiana (Barr et al., 1991). As is the case for all surveillance studies, prevalence can vary greatly depending on the diagnostic assay used to determine infection status and the geographic location.
Experimental infection studies of several wildlife species from the United States have confirmed the susceptibility of these animals to T. cruzi. In 1971, Virginia opossums developed patent infections after ingesting Rhodnius prolixis infected with the Tulahuen strain (Yaeger, 1971). However, neither Tulahuen strain nor R. prolixis are indigenous to the United States. In another study, four skunks were experimentally infected with a domestic dog isolate of T. cruzi (Davis et al., 1980). All animals developed patent infections with limited clinical signs. Other studies with US wildlife reservoirs included those conducted with wood rats that became infected with T. cruzi after ingesting infected
Triatoma protracta (Ryckman and Olsen, 1965). A raccoon T. cruzi isolate from
Maryland produced nonfatal infections in experimentally infected raccoons, young opossums (Didelphis marsupialis), and an adult primate (Macaca irus) (Walton et al., 19
1958), but not deer mice (Peromyscus leucopus). The same isolate produced similar results in experimentally infected pigs, lambs, kids, and calves (Diamond and Rubin,
1958). Infected animals had low levels of circulating parasites and some had few numbers of pseudocysts in heart tissue.
While different transmission routes for T. cruzi have been identified experimentally, the mechanism by which wildlife reservoirs predominately become infected with T. cruzi in the United States is unknown. Classic stercorarian vector transmission is unlikely because the two main reservoirs, raccoons and opossums, rarely use permanent dens, competent vectors are rarely found in or around temporary dens
(Walton et al., 1958), and native vectors, such as Triatoma sanguisuga, have delayed defecation times following the acquisition of a blood meal (Zeledón 1974). Vertical or transplacental transmission has been demonstrated several times in rodent models
(Andrade, 1982; Moreno et al., 2003) and naturally in humans (Hoff et al., 1978, Muños et al., 2007) but experimental studies in opossums disprove this method of transmission for the marsupial (Jansen et al., 1994).
Experimental per os infection trials in Virginia opossums (Yeager, 1971) and striped skunks (Davis et al., 1980) have implied direct oral transmission as the presumptive natural route with the ingestion of infected triatomid bugs or oral lavage with infected intestinal contents, respectively. Conversely, microcosm experiments have demonstrated that opossums rarely, if ever, predate infected bugs in simulated dens but acquire T. cruzi (Schweigmann et al., 1995). In addition to the ingestion of vectors, numerous claims have been made about the importance of carnivory in maintenance of 20 the sylvatic cycle (see Dias, 2006; Miles, 2004; Coura, 2006). However, no experimental data in natural reservoirs has been produced to support or refute such declarations.
Trypanosoma cruzi diagnostics
During the acute stage of infection there are a high number of parasites circulating in the blood, allowing for easy diagnosis by visualizing trypomastigotes in the blood.
This can be achieved by Giemsa or Wright‘s stain of thin blood smear, hemoculture, or xenodiagnosis. Xenodiagnosis is the feeding of uninfected, laboratory-raised bugs on the patient or host and examining the bugs for infection after 2 weeks. After the acute stage, parasitemias drop and diagnosis becomes more difficult. Multiple tests are often used to minimize false positives or negatives. Serology followed by one of the abovementioned techniques or animal inoculation is used during the chronic stage of Chagas‘ disease. In animals, the same methods may be used for diagnosis, but limitations exist because of low parasitemias and invalidation of rapid serological tests only evaluated in humans or dogs (Luquetti et al., 2003; Ponce et al., 2005; Cardinal et al., 2006).
Trypanosoma cruzi molecular evolution and genotyping
Prior to advances in molecular biology and genetics, differences in T. cruzi were based solely on growth characteristic and manifestations of disease in various hosts
(Hoare, 1972). These experiments suggested a heterogeneous population of T. cruzi
(Miles, 1979) and with the use of enzyme electrophoresis, heterogeneity was clearly demonstrated using isolates from wild and domestic mammals, humans, and vectors in
Brazil (Miles et al., 1978). The first segregation of genotypes was suggested when 21 isolates from opossums and a sylvatic triatomine species comprised a single group (type
I) that significantly differed from other isolates (type II) from humans, cats, rats, guinea pigs, house mice, and a domiciliary triatomine species. The differences in isolates from this single small area emphasized that little was known or understood of T. cruzi diversity and evolution.
Within the last 20 to 30 years, knowledge of the proliferative nature of T. cruzi and molecular epidemiology has yielded new perspectives on T. cruzi evolution. First, the organism was determined to be aneuploid (Gaunt et al., 2003), rather than the proposed and accepted diploid (Castro et al., 1981). As with many microbes, T. cruzi was considered a clonally proliferative parasite. With the continued use of isoenzyme electrophoresis, the classification was extended and the population was suggested to be comprised of multiple clones or groups (Tibayrenc et al., 1986). Not until the last decade has the understanding of T. cruzi population genetics been so dramatically changed. The theory of a solely clonal structure has been disregarded because of recent evidence for genetic exchange events (Gaunt et al., 2003; Machado and Ayala, 2001), thus changing the analyses used for phylogenetics from based on the Hardy-Weinberg principle to recombination tests (Tibayrenc, 2003).
Investigations of T. cruzi molecular evolution have become more complex and the analysis more rigorous. DNA sequencing techniques developed to determine phylogenetic lineages in a more high-throughput method are more useful and time-saving than the previously used isoenzyme electrophoresis. 22
One of the first successful analyses found two major lineages (Souto et al., 1996) similar to the findings of Miles et al (1978). Two lineages were identified within 88
South American T. cruzi isolates using the mini-exon and 24Sα rDNA gene targets.
Random amplified polymorphic DNA markers identified six phylogenetic lineages, TcI and TcII, with TcII having 5 subtypes (a-e) (Brisse et al., 2000). A more streamlined method (Figure 2.3) for typing T. cruzi isolates was developed where a series of PCR reactions for the mini- exon, 24Sα rRNA, and
18S rRNA are performed and the isolates typed according to the band patterns observed (Brisse et al.,
2001). Figure 2.3. Classic typing scheme of T. cruzi (Brisse et
al., 2001).
All six T. cruzi genotypes have been characterized among South American isolates from various host species (Yeo et al., 2005). Contrastingly, strains from Mexico and Central America (Guatemala) have been characterized as TcI and TcIIa, with a clear predominance of TcI isolates (Espinoza et al., 1998; Bosseno et al., 2002; Sánchez-
Guillén et al., 2006; Iwagami et al., 2007).
It has been previously suggested that the divergence of TcI and TcII lineages occurred approximately 80 million years ago (MYA) at the time of the geographical isolation of North and Central America from South America (Briones et al., 1999). 23
Clustering of all TcI sequences in previous phylogenetic studies occurs and may be due to a single origin of these strains (Iwagami et al., 2007; Subileau et al., 2009). Although genetic variability among TcI isolates from these studies is minimal, other studies have differentiated TcI isolates using the mini-exon and cytochrome b genes, sometimes suggesting the subdivision of the lineage similar to the subdivision of TcII (Herrera et al.,
2007; O‘Connor et al., 2007; Spotorno et al., 2008). Additionally, TcIIa isolates from
North America have been shown to cluster separately from South America isolates in previous studies (Iwagami et al., 2007; Subileau et al, 2009). However, these studies had limited to no isolates from the United States; therefore, the phylogeny of US T. cruzi is largely unknown.
Additional molecular studies have been completed to observe hybridization and genetic exchange in the T. cruzi population. Looking at ten intergenic regions of the T. cruzi genome in well-characterized isolates, TcIId and TcIIe have been confirmed as hybrid subgroups and provide evidence of genetic exchange from the parental groups TcI and TcIIb (Sturm et al., 2003). Another study compared nuclear and mitochondrial gene phylogenies to identify exchange events between lineages (Machado and Ayala, 2001).
While the majority of isolates were from South and Central America, three isolates were from the US, of which one exhibited genetic exchange. Experimental evidence of genetic exchange was revealed with the hybridization of clones (Gaunt et al, 2003); however, such events in nature are rare (Tibayrenc and Ayala, 2002), and their role in driving the evolution of the species has not been explored (Subileau et al., 2009). 24
Wildlife reservoirs and the genotype-strain dichotomy
Carlos Chagas first demonstrated that animals were natural hosts for T. cruzi when he successfully infected several lab animals, including marmosets, via the natural route using infected bugs (Chagas, 1909). He later went on to identify the first sylvatic reservoir, the nine-banded armadillo (Chagas, 1912). Since the early discoveries of
Chagas‘ disease, about 200 species or subspecies of wildlife infected with T. cruzi have been identified (Barretto and Ribeiro, 1979). For a species to be defined as a competent
T. cruzi reservoir, it must be important in the maintenance and transmission of the parasite. Animals considered reservoirs for T. cruzi are usually categorized according to the transmission cycle to which they contribute the most (e.g., sylvatic, peridomestic, and domestic cycles).
In Central and South America, armadillos and opossums are considered important sylvatic reservoirs. Reservoirs in the domestic and peridomestic cycles include rodents for both cycles, and opossums, including Philander frenata and Didelphis marsupialis, for peridomestic cycles. Mott et al (1978) suggested domestic dogs as important reservoirs for the domestic cycle. Although domestic dogs can develop cardiomyopathies that eventually lead to mortality, they live long enough for circulating parasites to be taken up by reduviids to perpetuate the life cycle. Because of their close proximity to humans, domestic dogs may be important reservoirs in the domestic and peridomestic cycles. In fact, in Argentina, risk of a human developing Chagas‘ disease is four times higher if a domestic dog sleeps in the bedroom (Crisante et al., 2006).
Although the presence of T. cruzi in wildlife has been confirmed, the use of isolates from these animals in molecular epidemiology studies has been limited. The first 25 was performed with 18 isolates from Georgia that were genotyped using riboprinting
(Clark and Pung, 1994). Another study performed by Barnabé et al, 2001 included 30 isolates; however, 18 isolates were from Clarke and Pung (1994). Associations between host species and parasite genotype have been previously suggested and are important in understanding both the domestic and sylvatic cycles of T. cruzi (Clark and Pung, 1994;
Briones et al., 1999; Yeo et al., 2005). A recent analysis of T. cruzi hosts throughout the
Americas indicated at least 48 hosts representing 17 families were infected with any of the six strains or lineages (Yeo et al., 2005). In that study, the South American Didelphis species were most commonly infected with type I, while armadillo species were infected with all the different types. Raccoons from the United States were primarily infected with
T. cruzi IIa. This trend in host-genotype preference suggests marsupials, such as the opossum (Didelphis spp.), are predominantly infected with type I, while raccoons
(Procyon lotor) are predominately infected with T. cruzi IIa; known exceptions include one raccoon from Louisiana from a previous study (Barnabé et al., 2001).
26
Table 2.1. Wildlife species found to be naturally infected with Trypanosoma cruzi through culture or direct observation of the parasite or serologic detection of antibodies reactive to T. cruzi. Species Sample or Assay n Prevalence Location and/or Reference (%) state Raccoon (Procyon Blood culture 54 22.1 Southeast Georgia Pung et al. 1995 lotor) Blood culture 30 43 St. Catherines Pietrzak and Pung Island, Georgia 1998 Blood smear 8 63 Tulsa, Oklahoma John and Hoppe 1986 Heart culture 47 2 Laurel, Maryland Herman and Bruce 2 1962 Blood culture 20 15 North Carolina Karsten et al. 1992 Heart and blood 35 14.3 East-central Olsen et al. 1964 culture Alabama Kidney culture 60 1.5 Southwest Georgia McKeever et al. 8 and northwest 1958 Florida Blood culture 33 12 Florida Schaffer et al. 1978 Blood culture 10 50 Georgia Schaffer et al. 1978 Blood culture 25 24 Texas Schaffer et al. 1978 Blood culture 5 N/A Maryland Walton et al. 1958 Blood culture 3 66 Tennessee Herwaldt et al., 2000 Indirect 22 47 South Carolina and Yabsley and Noblet Immunoflouresence 1 Georgia 2002 Assay Indirect 46 33 Fairfax County, Hancock et al. 2005 Immunoflouresence 4 Virginia Assay Indirect 9 0 South Texas Burkholder et al. Hemagglutination 1980 Assay Opossum Blood culture 39 15.4 Southeast Georgia Pung et al. 1995 (Didelphis virginiana) Heart and blood 12 13.5 Alabama Olsen et al. 1964 culture 6 Heart culture 21 0 Laurel, Maryland Herman and Bruce 9 1962 Blood culture 12 8.3 North Carolina Karsten et al. 1992 Blood culture 48 33.3 Southern Louisiana Barr et al. 1991b Kidney culture 55 16 Southwest Georgia McKeever et al. 2 and northwest 1958 Florida Nine-banded Blood culture 98 1.1 Southern Louisiana Barr et al. 1991 Armadillo (Dasypus novemcinctus) Blood culture 80 28.8 New Orleans, Yaeger 1988 Louisiana Direct 80 37.5 New Orleans, Yaeger 1988 Agglutination Louisiana
27
Species Sample or Assay n Prevalence Location and/or Reference (%) state Striped Skunk Kidney culture 30 1.0 Southwest Georgia McKeever et al. (Mephitis 6 and northwest 1958 mephitis) Florida Complement 1 100 Los Angeles, Ryan et al. 1985 Fixation and Direct California Agglutination Gray Fox Kidney culture 11 1.7 Southwest Georgia McKeever et al. (Urocyon 8 and northwest 1958 cinereoargenteus) Florida Indirect 26 8 South Carolina Rosypal et al. 2007 Immunofluorescent Antibody Test American Badger Indirect 8 25 South Texas Burkholder et al. (Taxidea taxus) Hemagglutination 1980 Assay Coyote (Canis Indirect 2 0 South Carolina Rosypal et al. 2007 rufus) Immunofluorescent Antibody Test Indirect 13 14.2 Central and Grogl et al. 1984 Immunoflouresence 4 southeast Texas Assay Indirect 15 12.8 South Texas Burkholder et al. Hemagglutination 6 1980 Assay Woodrat (Neotoma Blood smear 30 23.3 South Texas Burkholder et al. micropus) 1980 Pocket mouse Blood smear 25 16 South Texas Burkholder et al. (Perognathus 1980 hispidus) Mexican spiny Blood smear 11 9 South Texas Burkholder et al. pocket mouse 1980 (Liomys irrorattus) Grasshopper Blood smear 9 11.1 South Texas Burkholder et al. mouse 1980 (Onychomys leucogaster) Neotoma spp. and Blood smear 41 0.73 California Wood 1952 Peromyscus spp. 0 Neotoma spp. and Blood smear 86 4.7 Arizona Wood 1952 Peromyscus spp.
28
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43
CHAPTER 3
MOLECULAR TYPING OF TRYPANOSOMA CRUZI ISOLATES, UNITED STATES1
1 Roellig, D.M., Brown, E.L., Barnabé, C., Tibayrenc, M., Steurer, F.J., Yabsley, M.J. 2008. Emerging Infectious Diseases. 14: 1123-1125. Reprinted here with permission of publisher.
44
Abstract
Previous studies have characterized Trypanosoma cruzi from endemic regions.
With new human cases, increasing numbers of veterinary cases, and influx of potentially infected immigrants, understanding the ecology in the United States is imperative. We used a classical typing scheme to determine the lineage of 107 isolates from various hosts.
In Latin America, an estimated 10-12 million people are infected with
Trypanosoma cruzi (1), the etiologic agent of Chagas disease and an important contributor to heart disease within the region. Autochthonous human infections in the
United States have been reported in six individuals, with the most recent case reported from Louisiana (2). In addition, the parasite is euryxenous, meaning it is able to infect a broad range of hosts, including domestic dogs, woodrats, raccoons, opossums, armadillos, and non-human primates.
Associations between host species and parasite genotype have been previously suggested and are important in understanding both the domestic and sylvatic cycles of T. cruzi (3-5). Although previous studies conducted on US isolates suggest an association of
T. cruzi genotype with host, these studies were limited due to low sample numbers, low host diversity, and narrow geographic distribution (3, 5-8). In the current investigation, we have followed the molecular typing scheme proposed by Brisse et al. (9), in which isolates are delineated into one of the six lineages (Type I, IIa-IIe) based on size polymorphisms of several PCR markers. In doing this, we expand characterization of US isolates and reveal further evidence for correlations between biological characteristics, including host specificity, and genotype of T. cruzi. 45
The Study
We analyzed 107 isolates of T. cruzi from multiple species of free-ranging and captive wildlife, domestic animals, triatomine bug vectors, and humans who were autochthonously infected in the United States. Some isolates were obtained as liquid nitrogen-stored parasites from the Centers for Disease Control and Prevention, Pasteur
Institute, and the Southeastern Cooperative Wildlife Disease Study and were established in axenic LIT medium as previously described (10). Additional isolates were obtained from wild-trapped animals in axenic LIT medium or canine macrophage-cell culture as previously described (11). Isolated DNA was used as template for PCR amplification of three gene targets, mini-exon, D7 divergent domain of 24s alpha rRNA, and 18s rRNA, following methodologies previously published (9). The locality data and results of molecular typing of each isolate are shown in Table 1.
Only two genotypes were detected, T. cruzi I and T. cruzi IIa. The typical amplicon sizes of T. cruzi I and T. cruzi IIa isolates from the US are demonstrated in
Table 3.2. Atypical banding patterns and isolates that differ from the standard genotype from a particular host are also represented. With exception of the isolates from humans, a single primate, and a few raccoon isolates, placental mammal isolates, including those from raccoons, domestic dogs, ring-tailed lemurs, and skunks, were characterized as
Type IIa (Table 3.1). All remaining isolates, including those from the Virginia opossum
(Didelphis virginiana), triatomine vectors, humans, and rhesus macaques from the United
States were identified as Type I (Table 3.1). 46
Conclusions
In contrast to studies conducted on South American isolates where six genotypes of T. cruzi have been identified, only two genotypes (I and IIa) were identified in the current study. These data support investigations in Central America and Mexico that have found a paucity of genotypes (12, 13). Many investigations on T. cruzi evolutionary ecology reveal strict host-parasite specificity in regard to host species and parasite genotype (3-5), although exceptions have been observed. The presence of only two genotypes in the United States could be due to a lack of introduction of other genotypes or a lower diversity of natural reservoir hosts for T. cruzi in the United States compared to South America. A recent analysis of T. cruzi hosts in North and South America indicated at least 48 host species representing 17 families were infected with any of the six strains (5). Only six of these hosts have established populations in the United States, and US isolates from these species were only characterized as Types I or IIa (5).
Our data from US isolates corresponds with previous studies where Didelphis spp. are reservoirs for Type I T. cruzi (5); no infections with Type II parasites were observed.
The Virginia opossum is the only marsupial present in the United States, with its ancestors, which were possible hosts to T. cruzi I, migrating from South America approximately 4.5 million years ago (14). This evidence suggests that T. cruzi was not recently introduced to North America or the United States (6). Additionally, sufficient time may have passed for random and rare genetic exchange events to occur independent of those found in South American isolates (15), allowing the lineage to infect atypical reservoirs (i.e. raccoons) in North America. 47
The second major natural reservoir of T. cruzi in the United States is the raccoon.
In general, the non-primate placental mammals in our study were infected with Type IIa, a strain which is commonly found in sylvatic cycles in the southern cone of South
America. Our data confirms previous discrete typing of US isolates by MLEE and/or
RAPD analysis (6), in which eleven raccoons from Georgia were characterized as
Zymodeme 3 (equivalent to IIa). Although raccoons are predominately infected with T. cruzi IIa, four known exceptions include three isolates from Georgia and Florida in the current study and one raccoon from Louisiana from a previous study (6).These data are in contrast to typing data from Virginia opossum isolates which are all T. cruzi I, suggesting that opossums primarily maintain persistent infections with T. cruzi I.
All characterized human isolates from autochthonous US T. cruzi cases are T. cruzi I. The genotype is predominantly responsible for Chagas disease north of the
Amazon basin and is part of the domiciliary cycle of the parasite. Our findings correspond with data from Mexico where T. cruzi I is the predominate strain detected in humans (12). It would be interesting to differentiate biological characteristics and polymorphisms, using additional gene targets, in human Type I isolates compared to opossum, triatomine vector, and rhesus macaque isolates from the United States.
Additionally, comparing these US isolates and Mexican reference strains to those from
South America may reveal why Type I typically infects humans in North America while multiple strains are found in humans from South America.
Our current results provide additional support that T. cruzi has distinct genotypes that preferentially infect one host species or a group of hosts. T. cruzi types from humans and marsupials in the US are typically Type I, while raccoons, skunks, domestic dogs, 48 and prosimians are typically infected with Type IIa. Although we only detected T. cruzi I in triatomid bugs, previous studies have detected T. cruzi IIa in triatomids from the US
(6). Presently, the mechanism by which certain hosts develop persistent infections with a particular genotype of T. cruzi is unknown. Further analysis of isolates from an increased host diversity and geographic range should be pursued and determining basic infection dynamics of reservoir hosts experimentally infected with various T. cruzi genotypes may provide additional insight into the host-parasite dichotomy currently observed.
Acknowledgments
The authors thank B. Wilcox and B. Hanson (SCWDS), and D. Kavanaugh (USDA,
Wildlife Services) for field assistance. C. Paddock (CDC) kindly provided one isolate used in the study. P. Dorn (Loyola, LA) kindly provided blood for the isolation of one isolate. All animals used in this study were cared for in accordance with the guidelines of the Institutional Animal Care and Use Committee, and under animal use protocol approved by the Institutional Animal Care and Use Committee at the University of
Georgia. This study was supported by the National Institutes of Health, National Institute of Allergy and Infectious Diseases grant R15 AI067304.
Biographical Sketch
Ms. Roellig is a PhD student of Infectious Diseases at the University of Georgia. Her research interests are vector borne zoonotic diseases including Chagas disease in wildlife and tick-borne rickettsial pathogens. 49
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Table 3.1. Origin and lineage identification of the 107 United States isolates used in the study.
Host Isolate Origin Lineage Host Isolate Origin Lineage Human CA R California I Raccoon GA Rac 111 Ossabaw Island, GA IIa Corpus Christia Corpus Christi, TX I GA Rac 121 Ossabaw Island, GA IIa LC T cruzi New Orleans, LA I GA Rac 124 Ossabaw Island, GA IIa TC Californiaa Lake Don Pedro, CA I GA Rac 134 Whitehall Forest, GA IIa TX D Alano, TX I GA Rac 135 Whitehall Forest , GA IIa Domestic Dog Caesar Dog Not known IIa GA Rac 137 Whitehall Forest , GA IIa Dog Theisa, b, e Not known IIa GA Rac 141 Whitehall Forest , GA IIa Griffin Dog Hillsboro, TN I/IIa GA Rac 142 Whitehall Forest , GA IIa OK Dog Bartlesville, OK IIa GA Rac 143 Athens, GA IIa Samantha Dog South Carolina IIa GA Rac 144 Athens, GA IIa Smokey South Carolina IIa GA Rac 147 Woodbine, GA IIa USA Dog Ya California IIa GA Rac 148 Woodbine, GA IIa VA Opossum 92101601Pa Statesboro, GA I GA Rac 186 White Hall, GA IIa 93041401P cl1a Statesboro, GA I GA Rac 2 Ludiwici, GA I 93070103P cl2a Fort Stewart, GA I GA Rac 206 Athens, GA IIa FL Opo 15 Maclay State Park, FL I GA Rac 208 White Hall, GA IIa FL Opo 17 Wakulla Springs, FL I GA Rac 22 Victoria Bryant SP, GA IIa FL Opo 18 Wakulla Springs, FL I GA Rac 3 Athens, GA IIa FL Opo 2 Wakulla Springs, FL I GA Rac 45 Skidaway Island, GA IIa FL Opo 3 Wakulla Springs, FL I GA Rac 46 Skidaway Island, GA IIa FL Opo 717 Tampa, FL I GA Rac 51 Skidaway Island, GA IIa GA Opo 43 Chatham County, GA I GA Rac 52 Skidaway Island, GA IIa GA Opo 75 White Hall, GA I GA Rac 55 Skidaway Island, GA IIa Opossum 1970a New Orleans, LA I GA Rac 57 Skidaway Island, GA IIa USA Opossuma South Louisiana I GA Rac 61 Skidaway Island, GA IIa AU8 Auburn, AL I GA Rac 67 Athens, GA IIa FH4 South Georgia I GA Rac 68 Athens, GA IIa Raccoon 92122102Ra Statesboro, GA IIa GA Rac 69 Athens, GA IIa 93040701R cl1a Statesboro, GA IIa Maryland Rac Laurel, MD IIa 93053102R cl4a Harrold Preserve, GA IIa STC 10R cl3a St. Catherine‘s Island, GA IIa 93053103R cl3 Harrold Preserve, GA I STC 16R cl1a St. Catherine‘s Island, GA IIa 93071502R cl2a Fort Stewart, GA IIa STC 33R St. Catherine‘s Island, GA IIa 93072805R cl3a Fort Stewart, GA IIa STC 35R St. Catherine‘s Island, GA IIa 53
Host Isolate Origin Lineage Host Isolate Origin Lineage FL Rac 13 Maclay State Park, FL I/IIa STC 39R St. Catherine‘s Island, GA IIa FL Rac 14 Wakulla Springs, FL IIa STC 54R St. Catherine‘s Island, GA IIa FL Rac 15 Wakulla Springs, FL IIa STC 9R cl4a St. Catherine‘s Island, GA IIa FL Rac 26 Wakulla Springs, FL IIa TN Rac 18 Rutherford Co., TN IIa FL Rac 30 Wakulla Springs, FL IIa T. sanguisuga Floridaa Gainesville, FL I FL Rac 38 Maclay State Park, FL IIa Florida C16d Gainesville, FL I FL Rac 4 PAD Tallahassee, FL IIa Florida C1F8 Gainesville, FL I FL Rac 40 Wakulla Springs, FL IIa T. sang 5 cl1a Bulloch Co., GA I FL Rac 42 Wakulla Springs, FL IIa T. gerstackeri Triatoma 2 Texas I/IIa FL Rac 46 Tall Timbers, FL IIa Triatoma 3 Texas I FL Rac 48 Maclay State Park, FL IIa TxTg2 Texas I FL Rac 5 Torreya State Park, FL IIa RT Lemur Nilda St. Catherine‘s Island, GA IIa FL Rac 50 Wakulla Springs, FL IIa Clarence St. Catherine‘s Island, GA IIa FL Rac 51 Wakulla Springs, FL IIa Meg St. Catherine‘s Island, GA IIa FL Rac 7 Lake Talquin, FL IIa Rh. Macaque Monk RH89-40 Atlanta, GA (CDC) I/IIa FL Rac 9 Torreya State Park, FL IIa Texas Theisa Not Known I FR36c Pickens Co., SC IIa Nb Armadillo Armadillo 1973a New Orleans, LA I GA Rac 103 Ossabaw Island, GA IIa GA Arm 20 Ossabaw Island, GA IIa GA Rac 104 Ossabaw Island, GA IIa USA Armadilloa South Louisiana I GA Rac 107 Ossabaw Island, GA IIa Str. Skunk GA Sk 1 Ludiwici, GA IIa GA Rac 108 Ossabaw Island, GA IIa Table abbreviations: VA opossum= Virginia Opossum; RT lemur= Ring-tailed Lemur; Rh. Macaque= Rhesus Macaque; Nb Armadillo= Nine-banded Armadillo; Str. Skunk= Striped Skunk apreviously characterized by MLEE and/or RAPD analysis in 8. bpreviously characterized using microsatellite, 24S alpha rRNA, and COII genetic analysis in 16. cpreviously characterized by RAPD and mini-exon amplification in 8. dpreviously characterized (unspecified method) in 15. epreviously characterized by RAPD and MLEE analysis in 17. 54
Table 3.2. Approximate amplicon sizes of gene targets and lineage determination. Strain Mini-exon 24s alpha rRNA 18S rRNA Lineage FL Opo 15a 350 bp 110 bp 175 bp I GA Rac 103a None 120 bp 155 bp IIa FL Rac 5a 400 bp 120 bp 155 bp IIa 93053103R cl3 350 bp 110bp 175 bp I FL Rac 13 350 bp 110 bp and 120 bp 155 bp and 175 bp I/IIab FL Rac 46 400 bp 110 bp and 120 bp 155 bp I/IIab Griffin Dog 350 bp 110 bp and 120 bp 155 bp I/IIab Monk RH89-40 None 110 bp 155 bp I/IIab adenotes isolates used as representative banding patterns seen for classical lineage typing bI/IIa= atypical banding patterns, could not result in a clear definition of isolate as Type I vs Type IIa
55
CHAPTER 4
GENETIC VARIATION AND EXCHANGE IN TRYPANOSOMA CRUZI ISOLATES
FROM THE UNTED STATES1
1 Roellig, D.M., Savage, M.Y., Fujita, A.W., Barnabé, C., Tibayrenc, M., Steurer, F.J., Yabsley, M.J. To be submitted to Infection, Genetics, and Evolution. 56
Note: Nucleotide sequences reported in this paper have been deposited in the GenBank database (accession numbers: GU212879—GU213035).
Trypanosoma cruzi, the causative agent of Chagas disease, is a multiclonal parasite with high levels of genetic diversity and broad host and geographic ranges. Molecular characterization of South American isolates of T. cruzi has demonstrated homologous recombination and nuclear hybridization, as well as the presence of two main phylogenetic lineages (Type I and II). Few studies have extensively investigated such exchange events and genetic diversity in North American isolates. In the current study, we genetically characterized over 50 US isolates from wildlife reservoirs (e.g., raccoons, opossums, armadillos, skunks), domestic dogs, humans, nonhuman primates, and reduviid vectors from nine states (TX, CA, OK, SC, FL, GA, MD, LA, TN). Single nucleotide polymorphisms were identified in sequences of the mismatch-repair class 2
(MSH2) and Tc52 genes. Typing based on the two genes often paralleled genotyping by classic methodologies using mini-exon and 18S and 24Sα rRNA genes. Genetic exchange was determined by comparing sequence phylogenies of nuclear and mitochondrial gene targets, dihydrofolate reductase-thymidylate synthase (DHFR-TS) and the cytochrome oxidase subunit II- NADH dehydrogenase subunit I region (COII-ND1), respectively. We observed genetic exchange in several US isolates as demonstrated by incongruent mitochondrial and nuclear genes phylogenies, which confirms a previous finding of a single genetic exchange event in a Florida isolate. The presence of SNPs and evidence of genetic exchange illustrates that strains from the US are genetically diverse, even though only two phylogenetic lineages have been identified in this region. 57
Key words: Trypanosoma cruzi, trypanosome, genetic exchange, heterogeneity, genotype, Chagas‘ disease, evolutionary ecology
Introduction
Trypanosoma cruzi, the causative agent of Chagas‘ disease, is a clonally proliferative parasite with a heterogeneous population structure (Miles et al., 1978; Miles,
1979). It is a diverse parasite in its biological, molecular, and biochemical properties and has been detected in over 200 mammalian species, including humans (Barretto and
Ribeiro, 1979). Prior to advances in molecular biology and genetics, differences in T. cruzi were based solely on growth characteristics and manifestations of disease in various hosts (Hoare, 1972). Today, T. cruzi is segregated into two major discrete typing units,
TcI and TcII, with TcII having five subtypes (a-e) (Brisse et al, 2000; Brisse et al., 2001).
Characterizing a strain of T. cruzi into one of these six genotypes is useful in determining the evolutionary ecology of the parasite in a region, as well as, associating biological characters with disease manifestations. Also, the theory of a solely clonal population structure has been disregarded because of recent evidence for genetic exchange events
(Gaunt et al, 2003; Machado and Ayala, 2001).
Looking at ten intergenic regions of the T. cruzi genome in well-characterized isolates, TcIId and TcIIe have been confirmed as hybrid subgroups and provide evidence of genetic exchange from the parental groups TcI and TcIIb (Sturm et al., 2003).
Machado and Ayala (2001) have also illustrated genetic exchange by comparing nuclear and mitochondrial gene phylogenies and while the majority of isolates included in their study were from South and Central America, three isolates were from the US, of which 58 one exhibited genetic exchange. Additionally, experimental evidence of genetic exchange in a laboratory system was revealed with the hybridization of clones (Gaunt et al, 2003); however, such events in nature are rare (Tibayrenc and Ayala, 2002). Identifying genetic exchange may reveal clues as to the molecular evolution of T. cruzi, and strains identified as exhibiting this character may exhibit different biological characteristics compared with other strains of the same genotype.
Because T. cruzi is such a significant cause of morbidity and mortality in Central and South America, considerable research to charactertize T. cruzi has been conducted in these regions, but because human cases in the US are rare, little work has been conducted to characterize US isolates. Since 1955, six autochthonous human cases have been reported in the United States with the most recent occurring in 2006 (Woody and Woody,
1955; Schiffler et al, 1984; Ochs et al, 1996; Herwaldt et al, 2001; Dorn et al, 2007). In addition to these six cases, over 1,000 individual blood donors currently residing in the
United States were positive for antibodies reactive to T. cruzi (AABB, 2009). The objective of this study was to explore the molecular diversity of T. cruzi from the US.
Our goals were to obtain evidence of genetic exchange in US isolates and compare sequences of several gene targets with those from South America to understand the molecular evolution of the parasite in the United States. To accomplish these goals, nucleotide sequences of two nuclear genes, the mismatch-repair class 2 gene (MSH2) and the thiol-disulfide oxido-reductase Tc52 gene (Tc52), were compared with a selection of
T. cruzi isolates from the United States to identify single nucleotide polymorphisms that indicate heterogeneity and potential virulence differences. To investigate potential genetic exchange, the phylogenies of a nuclear gene [dihydrofolate reductase-thymidylate 59 synthase (DHFR-TS)] and mitochondrial gene targets [cytochrome oxidase subunit II-
NADH dehydrogenase subunit I region (COII-ND1)] were compared.
Materials and Methods
Isolates
T. cruzi was isolated from multiple species of free-ranging and captive wildlife, domestic animals, triatomine bug vectors, and humans who were autochthonously infected in the
United States; host and origin of each isolate is presented in Table 4.1. Some isolates were obtained as liquid nitrogen-stored parasites from the Centers for Disease Control and Prevention, Pasteur Institute, and the Southeastern Cooperative Wildlife Disease
Study and were established in axenic LIT medium as previously described (Castellani et al., 1967). Additional isolates were obtained from wild-trapped animals in axenic LIT medium or canine macrophage-cell culture as previously described (Brown et al., in press).
Molecular Technique
Template for polymerase chain reactions was obtained by boiling parasites for 15 min and using the resulting supernatant for DNA extraction with the DNeasy blood and tissue kit (Qiagen, Inc., Valencia, CA) following the manufacturer‘s protocol. PCR amplification was completed for the four gene targets, MSH2, Tc52, DHFR-TS, and
COII-ND1, following the respective previously published protocols (Augusto-Pinto et al.,
2003; Oury et al., 2005; Machado and Ayala, 2001). DNA extraction, amplification, and product analysis were performed in separate dedicated laboratory areas. A negative water control was included in each set of extractions and PCR reactions as contamination 60 controls. Sequencing reactions were performed at the Clemson University Genomics
Institute (Clemson, SC). Reactions were carried out with purified PCR product and amplicons were bidirectionally sequenced on an ABI 3100 Automated Sequencer
(Applied Biosystems, Foster City, CA). In the case of COII-ND1 products, reactions that did not yield complete sequences were identified and a heminested PCR reaction was performed using primers ND1.3A and COII.2A in the primary reaction and ND1.3S and
COII.A400 or COII.2A and COII.A400R in the secondary reactions (Machado and
Ayala, 2001).
Phylogenetic Analysis and Genotyping
Contiguous sequences were assembled in Sequencher and sequences aligned by the
Clustal Wallis method in Mega4. Three phylogenetic trees were created by neighbor- joining, minimum evolution and maximum parsimony methods from the alignment of each gene target with the bootstrap consensus tree being inferred from 500 replicates. The bat trypanosome T. cruzi marinkellei [593 (B3)] or T. brucei [TReu927] were used as the outgroups (Saitou and Nei, 1987; Eck and Dayhoff, 1966; Rzhetsky and Nei, 1992;
Felsenstein, 1985). A consensus tree was interpreted from the three methods.
Evolutionary distances were computed using the Kimura 2-parameter method (Kimura,
1980). Lineage typing of each isolate was performed with whole or partial sequences of the obtained gene sequence and a BLAST search was administered on GenBank to determine sequence identity with previously genotyped T. cruzi strains. 61
Results
Lineage typing of 51 T. cruzi isolates utilizing the four gene targets (Table 4.1) resulted in similar findings to genotyping using the mini-exon, D7 divergent domain of
24s alpha rRNA, and 18s rRNA PCR methodology (Brisse et al., 2001; Roellig et al.,
2008). Based on MSH2, Tc52, and DHFR-TS sequences, all human (Tc1), ring-tailed lemurs (TcIIa), armadillo (TcI or TcIIa) and skunk (TcIIa) isolates were genotyped as the same lineages previously determined (Roellig et al., 2008).
In contrast, some isolates from domestic dogs, Virginia opossums, raccoons, a rhesus macaque, and a Triatoma sanguisuga were classified as different lineages by different gene targets (Table 4.1). A raccoon isolate ‗FL Rac 13‘ that was previously thought to be a co-infection of TcI and TcIIa did not have evident polymorphisms (Tables
4.2 and 4.2). Sequences typed the isolate as TcI by the Tc52 and MSH2 genes, and it had
99% sequence identity to a TcIIa isolate for the DHFR-TS gene region. Another domestic dog isolate (USA Dog Y) was previously characterized as TcIIa but in this study was classified as TcI based on Tc52, MSH2, and DHFR-TS sequences; however mitochondrial sequences classified the strain as TcII. Lineage typing of raccoon isolate ‗GA Rac 143‘ indicated it as a TcIIa strain, but the Tc52 gene was 99% similar to a TcI isolate (Tep23).
The rhesus macaque isolate (Texas Theis) was previously genotyped as TcI, but for nuclear and maxicircle gene targets sequences obtained in this study it was 99-100% similar to TcIIa isolates. Seven Virginia opossum isolates demonstrated sequence similarity to the TcI genogroup of T. cruzi by nuclear gene analysis but were most similar to isolates in the TcII group when analyzing mitochondrial DNA sequences. 62
Single nucleotide polymorphisms (SNPs) were observed in the MSH2 and Tc52 genes of the analyzed sequences compared to TcI and TcIIa reference strains from South
America (Tables 4.2 and 4.3). The majority of sequences for the MSH2 (19 TcI and 24
TcIIa) and Tc52 (18 TcI/27 TcIIa) genes were identical (Tables 4.2 and 4.3). For the
MSH2 gene, only one or two nucleotides distinguished US TcI isolates from the reference strain while four to six nucleotides distinguished US TcIIa from the reference strain
(Table 4.2). For the Tc52 gene, three to four nucleotide substitutions distinguished the US
TcI isolates from the reference strain with one exception; a human isolate from a
California patient (Tc California) was identical to the South American reference strain
(Table 4.3). Numerous SNPs (13-22) distinguished the US and reference TcIIa strains.
Interestingly, the strain (TcIIa-TN Rac 18) that contained the most SNPs for both MSH2 and Tc52 genes was from a raccoon from Tennessee that was obtained during an investigation of an autochthonous case of Chagas‘ disease (Herwaldt et al., 2000).
Overall at least four SNPs were identified in these two genes that could be used to separate US isolates of TcI and TcIIa from the two South American reference strains
(Tables 4.2 and 4.3). Numerous other SNPs were identified but these occurred in only one or two isolates. For the Tc52 and MSH2 genes of US TcIIa isolates, three and four additional nucleotides, respectively, differed from the reference strain but these four nucleotides were the same as TcI isolates. Several nucleotide changes resulted in amino acid changes (Tables 4.2 and 4.3). The phylogenies of the two gene targets show the clustering of isolates with their respective genotype, including those that exhibited unique
SNPs (Figure 4.1). 63
Phylogenetic analysis of the nuclear gene region, DHFR-TS, supported the findings of the Tc52 and MSH2 gene analyses and resulted in a tree that had a similar topology to a previous study (Machado and Ayala, 2001) with four major clades, three of which represented TcII genotypes (Figure 4.2). Interestingly, the TcIIa isolates from the
United States were included in a well supported (66-75%) clade separate from South
American TcIIa isolates. Within the US TcIIa clade, a smaller clade including four isolates was present but this separation had lower bootstrap support (64-65%). Similar to sequence analysis results for the DHRF-TS gene of various TcI isolates (Machado and
Ayala, 2002), limited differences were noted within the US isolates as no separation of
TcI sequences was present (Figure 4.2). TcIIa isolates had 99% sequence identity to the
CANIII reference strains with 7 SNPs; TcI isolates had 99% sequence identity to the
Silvio X10 cl4 reference strain with 4 SNPs. The phylogeny for the COII-ND1 mitochondrial region showed greater divergence with four clades containing additional divisions (Figure 4.3). Eight isolates that were classified as TcI by analysis of various nuclear genes (e.g., 18S, mini-exon, 24S alpha, MSH2, Tc52, DHFR-TS) were classified as TcII by phylogenetic analysis of COII-NDI sequences. These eight isolates are highlighted in Figures 4.2 and 4.3.
Within TcIIa DHFR-TS sequences, two SNPs were identified with one being unique to isolate GA Rac 45; there were no SNPs among TcI DHFR-TS sequences (data not shown). Single nucleotide polymorphisms within TcIIa isolates were not noted, but compared with the CANIII reference strain, all TcIIa ND1 sequences had 96% sequence identity and 36 SNPs (data not shown). Paralleling differences between TcI isolates illustrated in the ND1 phylogeny, 88 SNPs were identified within TcI sequences so that 64 two sequence groups were found with all sequences within a group having the same sequence (data not shown). Interestingly, these two groups correspond to the TcI strains that exhibit genetic exchange and those that do not. Compared with the TcI reference strain, those that did exhibit exchange events had 92% sequence similarity to the Silvio
X10 cl4 reference strain, with 85 SNPs; sequences without genetic exchange had 98% sequence identity to Silvio X10 cl4, with 19 SNPs.
Discussion
In the current investigation, genetic diversity was demonstrated among T. cruzi isolates from the United States. T. cruzi strains are currently categorized into two major lineages, TcI and TcII, with TcII further subdivided into 5 genotypes (TcIIa-TcIIe). It has been previously suggested that the divergence of TcI and TcII lineages occurred approximately 80 MYA at the time of the geographical isolation of North and Central
America from South America, allowing isolation of placental and marsupial mammals of the two continents and independent evolution of T. cruzi infecting either group of mammals (Briones et al., 1999). All six genotypes have been characterized from South
American isolates from various host species (Yeo et al., 2005). Contrastingly, strains from Mexico and Central America (Guatemala) have been characterized as TcI and
TcIIa, with a clear predominance of TcI isolates (Espinoza et al., 1998; Bosseno et al.,
2002; Sánchez-Guillén et al., 2006; Iwagami et al., 2007). Isolates from the United
States, have also been characterized only as TcI or TcIIa (Clark and Pung, 1994; Barnabé et al, 2001; Roellig et al., 2008). Further confirming the paucity of genotypes in North
America, in the current study, sequences of additional gene targets had sequence identity 65 only to either TcI or TcIIa. Regardless of gene target, TcIIa isolates were clearly distinguished from the South American TcIIa reference strain which provides additional evidence for considerable divergence within this lineage (Machado and Ayala, 2001;
Westenberger et al., 2005)
To further investigate genetic diversity among US T. cruzi isolates, the sequences of two nuclear genes, Tc52 and MSH2, were analyzed to identify SNPs. Tc52 is a single- copy gene constitutively-expressed in all developmental stages of T .cruzi and is implemented in the immune response to T. cruzi infection, where it suppresses T-cell proliferation by scavenging cysteine and glutathione (GSH) (Ouaissi et al., 1995a;
Ouaissi et al., 1995b). Similar to previous findings (Oury et al., 2005; Mathieu-Daudé et al., 2007), numerous SNPs were found in the sequences of 53 isolates analyzed in this study. Of the 47 SNPs identified, 17 resulted in amino acid changes, several of which have been previously linked to GSH binding (Oury et al., 2005). Changes in amino acids resulting from these SNPs may account for differences in virulence betweens strains due to changes in GSH binding efficacy. Additional research is needed to determine if there is an association of these SNPs with biological differences, either between TcI and TcIIa or between isolates from the US and those from South America.
Polymorphisms were also identified in MSH2, a homologue of the mutS gene of other eukaryotes (Augusto-Pinto et al., 2001). The MSH2 protein is a part of the mismatch repair machinery that binds base-base mismatches and excises and repairs them. In T. cruzi, MSH2 is also a single copy gene that is constitutively-expressed in all life stages of the parasite (Augusto-Pinto et al., 2001). In the current investigation, we identified 21 SNPs of the MSH2 gene, including several that could distinguish between 66
TcI and TcIIa strains. Previous findings suggested that SNPs in TcII lineage had decreased mismatch-repair ability compared to TcI strains (Augusto-Pinto et al., 2003).
In our study, the majority of genetic variability was noted in the TcIIa isolates.
Interestingly, TcIIa isolates from the US tend to be less virulent to laboratory mice and to date, no human infections with this genotype have been reported in North America
(Roellig et al., unpublished). However, TcIIa strains have been isolated from primates, prosimians, and domestic dogs (Pung et al., 1998; Hall et al., 2007; Roellig et al., 2008).
Heterogeneity based on sequence differences, therefore, may lead to differences in genome maintenance, perhaps rendering those with decreased mismatch-repair efficiency less virulent.
In addition to identifying sequence differences in these US isolates, phylogenies were constructed for DHFR-TS (nuclear) and COII-ND1 (mitochondrial) to elucidate genealogical relationships among isolates and illustrate evidence for genetic exchange.
The nuclear phylogeny of DHFR-TS exhibited four major clades, three consisting of TcII strains and one TcI. Isolates of TcI from the US clustered with S. American isolates, illustrating the limited genetic variability of the lineage reported in previous studies with this gene (Machado and Ayala, 2001; Iwagami et al., 2007). Although genetic variability among TcI isolates was minimal for nuclear gene targets in this study, considerable biological differences between isolates have been previously noted (Barrera-Pérez et al.,
2001; Bértoli et al., 2006; Lisboa et al., 2007). Other studies have differentiated TcI isolates using the mini-exon and cytochrome b genes, sometimes suggesting the subdivision of the lineage similar to the subdivision of TcII (Herrera et al., 2007;
O‘Connor et al., 2007; Spotorno et al., 2008). More variation was noted for TcII isolates 67 with the US TcII clade being separate from the S. American TcIIa isolates. This division between N. and S. American TcIIa strains may be evidence of the independent evolution of N. American TcIIa strains from its ancestral S. American strains, as was suggested previously (Briones et al., 1999) and is supported by sequence analysis of the MSH2 and
Tc52 genes and phylogenetic analysis of the COII-ND1 gene.
The phylogeny of COII-ND1 demonstrated greater genetic diversity with additional clustering occurring within the four clades present. As with the nuclear phylogeny, TcIIa strains from the US diverged from S. American TcII strains. Additional clusters within the US TcIIa clade indicate additional genetic diversity within the group; however, several of these subclades had low bootstrap support. The clade representing
TcI strains contained both US isolates from this study and South American isolates, which is consistent with the nuclear DHFR-TS phylogeny. As previously suggested, the clustering of all TcI sequences may be due to a single origin of these strains (Iwagami et al., 2007; Subileau et al., 2009). It is also possible that TcI represents a more recent introduction or spread into North America compared with TcIIa, which may have been separate from the South American strains for a significant period of time which would allow divergence.
The most compelling finding from the COII-ND1 phylogeny is the clustering of several TcI strains (classified based on several nuclear genes) within the US TcIIa clade.
Incongruencies between nuclear and mitochondrial phylogenies have been previously reported with S. American isolates and a single US isolate and is interpreted as evidence of rare genetic exchange events in the T. cruzi population (Machado and Ayala, 2001;
Brisse et al., 2003). While T. cruzi is accepted as a predominately clonally propagating 68 population, these findings in addition to in vitro demonstration of genetic recombination illustrates that genetic exchange does occur, albeit rarely (Machado and Ayala, 2001;
Brisse et al., 2003; Gaunt et al., 2003). In the current study, several TcI sequences represent isolates that may have undergone genetic exchange in comparison to no TcII sequences. This suggests that TcI isolates may be more susceptible or likely to have recombination (Subileau et al., 2009). While these phylogenies can be associated with genetic exchange, the role of such events in driving the evolution of the species has not been explored (Subileau et al., 2009). In the current study, we identified several isolates with evidence of genetic exchange. While this region has only two (TcI and TcIIa) of the six genealogical lineages circulating in mammal populations, the presence of SNPs and evidence of genetic exchange suggest that parasite populations in the United States are genetically diverse. Associating these molecular characters with biological differences is the next step in investigating the diversity of T. cruzi in the region.
Acknowledgments
The authors thank E. Brown, B. Wilcox and B. Hanson (SCWDS), and D.
Kavanaugh (USDA, Wildlife Services) for field assistance. This study was supported by the National Institutes of Health, National Institute of Allergy and Infectious Diseases grant R15 AI067304.
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76
Table 4.1. Lineage typing of Trypanosoma cruzi isolates from the United States. Gene target Host Isolate Origin Lineageb Tc52 MSH2 DHFR-TS COII-ND1 Human TC CC Corpus Christi, TX I I I Ic n.d. CA R California I I I I I TC California Lake Don Pedro, TX I I I n.d. Ic Domestic Dog Caesar Dog Not known IIa IIa IIa IIa II Dog Theis Not known IIa IIa IIa IIa II OK Dog Bartlesville, OK IIa IIa IIa IIa II Samantha Dog South Carolina IIa IIa IIa IIa II Smokey South Carolina IIa IIa IIa IIa IIc USA Dog Y California IIa I I I II VA Opossum 92101601P cl2 Statesboro, GA n.d. I I I II 93041401P cl1 Statesboro, GA I I I I IIc 93070103P cl2 Fort Stewart, GA I I I I II FH4 South Georgia I I I I II FL Opo 2 Wakulla Springs, FL I I I I II FL Opo 3 Wakulla Springs, FL I I I I II FL Opo 15 Maclay State Park, FL I I I n.d. II Opossum 1970 New Orleans, LA I I I I Ic USA Opossum South Louisiana I I I I I Raccoon 92122102R Statesboro, GA IIa IIa IIa IIa II 93040701R cl2 Statesboro, GA IIa IIa IIa IIa II 93053103R cl3 Harrold Preserve, GA I I I I I 93071502R cl2 Fort Stewart, GA IIa IIa IIa IIa II 93072805R cl3 Fort Stewart, GA IIa IIa IIa IIa II FL Rac 13 Maclay State Park, FL I/IIa I I IIa n.d. FL Rac 15 Wakulla Springs, FL IIa IIa IIa IIa II FL Rac 30 Wakulla Springs, FL IIa IIa IIa IIa II FL Rac 46 Tall Timbers, FL IIa IIa IIa IIa n.d. FL Rac 5 Torreya State Park, FL IIa IIa IIa IIa n.d. FL Rac 7 Lake Talquin, FL IIa IIa IIa IIa II 77
Gene target Host Isolate Origin Lineageb Tc52 MSH2 DHFR-TS COII-ND1 FL Rac 9 Torreya State Park, FL IIa IIa IIa IIa II GA Rac 107 Ossabaw Island, GA IIa IIa IIa IIa II GA Rac 134 Whitehall Forest, GA IIa IIa IIa IIa II GA Rac 143 Athens, GA IIa I IIa IIa II GA Rac 45 Skidaway Island, GA IIa IIa IIa IIa n.d. GA Rac 69 Athens, GA IIa IIa IIa IIa II Maryland Rac Laurel, MD IIa IIa IIa IIa IIc STC 10R cl3 St. Catherine‘s Island, GA IIa IIa IIa IIa II STC 35R St. Catherine‘s Island, GA IIa IIa IIa IIa II TN Rac 18 Rutherford Co., TN IIa IIa IIa IIa II T. sanguisuga Florida Gainesville, FL I I I I n.d. Florida C1F8 Gainesville, FL I I I I II T. sang 5 cl1 Bulloch Co., GA I IIa IIa IIa n.d. RT lemur Nilda St. Catherine‘s Island, GA IIa IIa n.d. IIa n.d. Clarence St. Catherine‘s Island, GA IIa IIa IIa IIa IIc Meg St. Catherine‘s Island, GA IIa IIa IIa n.d. IIc Rh. Macaque Texas Theis Not known I IIa IIa n.d. II Nb Armadillo Armadillo 1973 New Orleans, LA I I I I I GA Arm 20 Ossabaw Island, GA IIa IIa IIa IIa II USA Armadillo South Louisiana I I I I I Str. Skunk GA Sk 1 Ludiwici, GA IIa IIa IIa n.d. IIc a Table abbreviations: VA opossum= Virginia Opossum; RT lemur= Ring-tailed Lemur; Rh. Macaque= Rhesus Macaque; Nb Armadillo= Nine-banded Armadillo; Str. Skunk= Striped Skunk; n.d.= not determined bpreviously characterized using mini-exon, D7 divergent domain of 24s alpha rRNA, and 18s rRNA genetic analysis in Roellig et al., 2008. cpartial sequences were analyzed. 78
Table 4.2. Nucleotide sequence variations within the MSH2 gene sequence of 49 T. cruzi isolates from the United States compared to reference strains. Genotype of each isolate precedes the isolate name. Nucleotide positions correspond to sites from SilvioX10 (Genbank AY540739). Dots represent nucleotide site identical to reference strain (either Silvio X10 for TcI or CanIII for TcIIa). Nucleotide Position
Genotype/Isolate 71 97 109 172 244 351 367 373 403 404 460 478 490 500 634 645 658 735 750 775 816
TcI Reference strain C A G A C T A G G G A T C A T C A C A G A (Silvio X10 cl1) TcI – 17 US sequences* • • • • • A • • • • • • • • • • • • • • • TcI-USA Opossum • • • • • A • • • • • • • • • • • • • • G TcI-93070103P cl2 • • • • • A • • • • • Y • • Y S • • T • • Amino acid change V→D S→C
TcIIa Reference strain G G G G T A G A G G A C T C C G A C A G A (CanIII) TcIIa-24 US sequences† • A • • • • • G • • T • C • • • • • • • • TcIIa-TN Rac 18 • A • • • • • G A A T • C M • • • • • • • TcIIa-STC 10R cl3 • A • • • • • G • • T • C • • • G • • • • TcIIa-Fl Rac 7 • A • • • • • G • • T • C • • • • G • • • TcIIa-GA Rac 107 • A • • • • • G • • T • C • • • • • • A • TcIIa-FL Rac 30 • A • • • • • G • • T • C • • • T • • • • TcIIa-T. sang 5 cl1 • A T • • • • G • • T • C • • • • • • • • Amino acid change V→M I→L T→R *The following 17 US TcI sequences were identical: Human isolates (TC CC, CA R, TC California), domestic dogs (USA Dog Y), Virginia opossums (92101601P cl2, 93041401P cl2, FH4, FL Opo 2, FL Opo 3, FL Opo 15, Opossum 1970), armadillos (Armadillo 1973, USA Armadillo), triatomine bugs (Florida C1F8, Florida), and raccoons (93053103R cl3, FL Rac 13).
†The following 24 US TcIIa sequences were identical: Raccoons (FL Rac 9, 92122102R, 93071502R cl2, 93040701R cl1, 93072805R cl3, FL Rac 15, FL Rac 46, FL Rac 5, GA Rac 134, GA Rac 143, GA Rac 45, GA Rac 69, Maryland Rac, STC 35R), domestic dogs (Samantha Dog, Caesar Dog, Dog Theis, OK Dog, Smokey), ring-tailed lemurs (Clarence, Meg), rhesus macaque (Texas Theis), striped skunk (GA Sk 1), and armadillo (GA Arm 20). 79
Table 4.3. Nucleotide sequence variations within the Tc52 gene sequence of 50 T. cruzi isolates from the United States compared to reference strains. Genotype of each isolate precedes the isolate name. Nucleotide positions correspond to sites from P209 (Genbank EF065175). Dots represent nucleotide site identical to reference strain (either P209 for TcI or CanIII for TcIIa). Dashes represent missing nucleotides. Nucleotide Position
Genotype/Isolate 8 9 1 1 151- 155- 159- 2 2 2 2 3 3 3 4 5 5 5 6 6 6 6 6 6 7 8 8 8 875- 9 967- 9 9 1 1 1 1 1 1 1 1 5 1 2 4 153 156 160 0 2 3 4 3 5 9 4 0 6 8 0 2 2 2 5 7 2 1 5 7 876 2 968 7 8 0 0 1 1 1 2 2 2 1 8 0 1 1 2 6 7 2 3 0 0 8 3 1 5 8 9 0 8 6 2 3 0 1 0 2 4 0 4 4 5 6 9 1 8 3 0 3 1 2 5 TcI reference strain C A T G ACA GG CT A A T T A C T G A G A A G T A A C C T G A CC A TA T C T T C G T A A A (P209) TcI – 17 US sequences* • • • • ••• •• •• • • • • • • • • • • • • • • • • • • • • • •• G •• • • • G • • • • G •
TcI-TC California • • • • ••• •• •• • • • • • • • • • • • • • • • • • • • • • •• • •• • • • • • • • • • •
TcI-93041401P cl1 • • C • ••• •• •• • • • • • • • • • • • • • • • • • • • • • •• G •• • • • G • • • • G •
TcI-FL Opo 2 • • • • ••• •• •• • • • • • • • • • • • • • • • • A • • • • •• G •• • • • G • • • • G •
Amino acid change V L→ T R → P → → A K M
TcIIa reference strain A A T G ACA GG CT G A T C G G C G G G G G A C G T A T G A G AT G CG C G A T G G A G G G (CanIII) TcIIa- 27 US sequences† C • • • ••• •• •• A G G T • • • A • • A A • • A • • • T • A C• • •• • • • • • A • • • •
TcIIa-TN Rac 18 C C • A GGT CA GA A G G T • • • A • • A A • • A • • • T • A C• • •• • • • • • A • • • •
TcIIa-FL Rac 7 C • • • ••• •• •• A G G T • • • A • A A A • • A • • • T • A C• • •• • • • • • A • • • •
TcI/IIa-Texas Theis • • • • ••• •• •• A G G • • • • A • • A A • • A • • • T • A C• • •• • • • • • A • • • •
Amino acid change E E Y K→Y E→ L→ S A A R G A A → → → K E → → → → → → → A A W A T T K W T T *The following 17 US TcI sequences were identical: Human isolates (TC CC, CA R), domestic dogs (USA Dog Y), Virginia opossums (USA Opossum, Opossum 1970, 92101601P cl2, FH4, 93070103P cl2, FL Opo 3, FL Opo 15), armadillos (Armadillo 1973, USA Armadillo), triatomine bugs (Florida C1F8, Florida), and raccoons (GA Rac 143, 93053103R cl3, FL Rac 13).
†The following 27 US TcIIa sequences were identical: Raccoons (STC 10R cl3, FL Rac 9, 92122102R, 93071502R cl2, 93040701R cl1, 93072805R cl3, FL Rac 15, FL Rac 46, FL Rac 5, FL Rac 30, GA Rac 134, GA Rac 45, GA Rac 69, GA Rac 107, Maryland Rac, STC 35R), domestic dogs (Samantha Dog, Caesar Dog, Dog Theis, OK Dog, Smokey), ring-tailed lemurs (Clarence, Meg, Nilda), striped skunk (GA Sk 1), triatomine bug (T sang5 cl1), and armadillo (GA Arm 20). 80
81
Figure 4.1. Evolutionary relationships among e mismatch-repair class 2 gene (MSH2) and the thiol-disulfide oxido-reductase Tc52 gene (Tc52) from 50 and 51 Trypanosoma cruzi isolates, respectively. Three phylogenetic trees were created by neighbor-joining (NJ), minimum evolution (ME), and maximum parsimony (MP) methods from the alignment of each gene target and a consensus tree was interpreted. Numbers at the branches are bootstrap values >50% (500 replicates) for the same nodes of the NJ, ME, MP trees. Evolutionary distances were computed using the Kimura 2-parameter method (Kimura, 1980). Italicized names are samples from the current study. ▲= the 17 US TcI isolates that were identical; ●= the 24 or 27 US TcIIa isolates that were identical. 82
83
Figure 4.2. Evolutionary relationships among dihydrofolate reductase-thymidylate synthase (DHFR-TS) from 43 Trypanosoma cruzi isolates. Three phylogenetic trees were created by neighbor-joining (NJ), minimum evolution (ME), and maximum parsimony (MP) methods from the alignment of each gene target and a consensus tree was interpreted. Numbers at the branches are bootstrap values >50% (500 replicates) for the same nodes of the NJ, ME, MP trees. Evolutionary distances were computed using the Kimura 2-parameter method (Kimura, 1980). Italicized names are samples from the current study. The ten isolates with positions incongruent to the mitochondrial phylogenies (Fig. 4.3) are highlighted. 84
85
Figure 4.3. Evolutionary relationships among cytochrome oxidase subunit II- NADH dehydrogenase subunit I region (COII-ND1) from 43 Trypanosoma cruzi isolates. Three phylogenetic trees were created by neighbor-joining (NJ), minimum evolution (ME), and maximum parsimony (MP) methods from the alignment of each gene target and a consensus tree was interpreted. Numbers at the branches are bootstrap values >50% (500 replicates) for the same nodes of the NJ, ME, MP trees. Evolutionary distances were computed using the Kimura 2-parameter method (Kimura, 1980). Italicized names are samples from the current study. The ten isolates with positions incongruent to the nuclear phylogenies (Fig. 4.2) are highlighted.
86
CHAPTER 5
GENETICALLY DIFFERENT ISOLATES OF TRYPANOSOMA CRUZI ELICIT
DIFFERENT INFECTION DYNAMICS IN RACCOONS (PROCYON LOTOR) AND
VIRGINIA OPOSSUMS (DIDELPHIS VIRGINIANA)1
1 Roellig, D.M., Ellis, A.E., Yabsley, M.J. 2009. International Journal for Parasitology. 39: 1603-1610. Reprinted here with permission of publisher. 87
Trypanosoma cruzi is a genetically and biologically diverse species. In the current study we determined T. cruzi (Tc) infection dynamics in two common North American reservoirs, Virginia opossums (Didelphis virginiana) and raccoons (Procyon lotor).
Based on previous molecular and culture data from naturally-exposed animals, we hypothesized that raccoons would have a longer patent period than opossums, and raccoons would be competent reservoirs for both TcI and TcIIa, while opossums would only serve as hosts for TcI. Individuals (n=2 or 3) of each species were inoculated with
1x106 culture-derived T. cruzi trypomastigotes of TcIIa [North America (NA)-raccoon],
TcI (NA- opossum), TcIIb (South America-human), or both TcI and TcIIa. Parasitemias in opossums gradually increased and declined rapidly; whereas, parasitemias peaked sooner in raccoons and they maintained relatively high parasitemia for 5 weeks.
Raccoons became infected with all three T. cruzi strains, while opossums only became infected with TcI and TcIIb. Although opossums were susceptible to TcIIb, infection dynamics were dramatically different compared with TcI. Opossums inoculated with
TcIIb seroconverted, but parasitemia duration was short and only detectable by PCR. In addition, raccoons seroconverted sooner [3-7 days post inoculation (DPI)] than opossums
(10 DPI). These data suggest that infection dynamics of various T. cruzi strains can differ considerably in different wildlife hosts.
KEYWORDS: Trypanosoma cruzi, experimental infection, raccoon, opossum, United
States, trypanosome
88
Introduction
Trypanosoma cruzi, the causative agent of Chagas‘ disease, has a wide host and geographic range. Approximately 200 species or subspecies of wildlife have been identified with T. cruzi infection (Barretto and Ribeiro, 1979) in a geographic range encompassing most of the Americas. Within the various host species that can become infected with T. cruzi, the genotype of the parasite may vary and an association between host and genotype has been strongly supported by molecular typing of isolates from autochthonously infected wild and domestic animals, humans, and vectors (Clark and
Pung, 1994; Briones et al., 1999; Yeo et al., 2005; Roellig et al., 2008). Although all six phylogenetic lineages, TcI and TcII (a-e), are found in South America, only TcI and
TcIIa have been identified in the United States (Clark and Pung, 1994; Barnabé et al.,
2001; Hall et al., 2007; Roellig et al., 2008).
Raccoons and Virginia opossums are considered important wildlife reservoirs in the United States with prevalence as high as 63% in raccoons (John and Hoppe, 1986) and 33% in Virginia opossums (Barr et al., 1991a). Lineage typing of T. cruzi from these two species has revealed a trend where the majority of T. cruzi isolates from raccoons have been TcIIa while all of the Virginia opossum T. cruzi isolates have been TcI
(Barnabé et al., 2001; Roellig et al., 2008). While there is evidence for a host-genotype dichotomy, the mechanisms driving the strain preference are unknown and experimental evidence of such a preference has not been demonstrated previously in these two wildlife reservoir species.
In addition to differences in T. cruzi genotypes isolated from naturally-infected raccoons and Virginia opossums, prevalence based on isolation success varies 89 considerably. Significantly more wild raccoons are culture positive compared to wild
Virginia opossums which suggests either differential exposure or infection dynamics
(Brown et al.,in press). The question of differential exposure was examined by Brown et al. (in press); sympatric opossums and raccoons from 10 counties in Georgia (USA) were tested for T. cruzi exposure and no difference in seroprevalence was noted between opossums and raccoons from the same area. Therefore, differences in prevalence between these two hosts based on culture isolation attempts are likely the result of differences in infection dynamics.
In the present study, the infection dynamics of the two major US reservoirs were determined after inoculation with different genotypes of T. cruzi. Our objectives were to determine if any differences in host susceptibility to different genotypes exists by measuring the duration and magnitude of parasitemias, time to seroconversion, presence of tissue stages, and histopathologic lesions. Based on previous genetic studies of field isolates, we hypothesized that raccoons would develop patent infections with TcI and
TcII strains, while Virginia opossums would develop patent infections with only TcI.
Additionally, because T. cruzi is more frequently isolated from raccoons compared to opossums, despite similar exposure rates (Brown et al., in press), we hypothesized that raccoons would develop higher parasitemias that would be maintained for longer periods compared with Virginia opossums.
90
Materials and methods
Inoculation material
The two North American T. cruzi isolates used in this study were originally isolated from a naturally-infected raccoon [FL-RAC9 (TcIIa)] and Virginia opossum [FL-
OPO3 (TcI)] from northwestern Florida (Roellig et al., 2008). One South American T. cruzi strain, Y (TcIIb), was generously provided by Dr. Rick Tarleton (University of
Georgia). The FL-OPO3 isolate was used as a representative TcI strain and the FL-
RAC9 and Y strains were used as representative TcII strains. Each strain was molecularly typed utilizing the mini-exon intergenic spacer gene, 24Sα rDNA D7 divergent domain, and size-variable domain of the 18S rRNA gene (Brisse et al., 2001;
Roellig et al., 2008).
Epimastigotes were passaged from Liver-Infusion Tryptose (LIT) medium into
DH82 canine macrophage monolayers at 1:5 dilutions to yield the infective culture- derived trypomastigotes. Trypomastigotes were pelleted from culture by centrifugation at 1620 x g for 15 min and resuspended in minimum essential medium (MEM). The concentration of parasites in suspension was determined with a hemocytometer.
Animals and experimental design
Ten juvenile raccoons obtained from Ruby Fur Farm, Inc. (New Sharon, IA) were housed individually or in pairs in climate-controlled animal housing at the College of
Veterinary Medicine, University of Georgia (Athens, GA). Fourteen juvenile Virginia opossums of two wild-trapped females from Athens, GA were housed with and reared by their respective mothers until weaning at approximately 12 weeks, after which they were individually housed, in climate-controlled animal housing at the College of Veterinary 91
Medicine, University of Georgia (Athens, GA). All animals used in this study were cared for in accordance with the guidelines of the Institutional Animal Care and use Committee and under an animal use protocol approved by this committee at the University of
Georgia. Animals were given food and water ad libitum. Before use, all raccoons and opossums were determined to be negative for antibodies reactive with T. cruzi (as described below). Both opossum mothers were also determined to be negative for T. cruzi by PCR, culture, and serology.
For both species, animals were randomly separated into four experimental groups and one negative control group. Raccoons (n=2) were inoculated intravenously (IV) and
Virginia opossums (n=3) were inoculated intraperitoneally (IP) with 1 x 106 culture- derived trypomastigotes of one of four inoculums: FL-OPO3 (TcI), FL-RAC9 (TcIIa), Y
(TcIIb), or equal parts FL-OPO3 strain and FL-RAC9 strain (5 x 105 of each). Negative controls (n=2) for both species were similarly inoculated with an equivalent volume of culture medium.
For handling and blood collection, raccoons were anaesthetized with an intramuscular (IM) injection of a mixture of 20 mg/kg ketamine (Fort Dodge
Laboratories, Inc., Fort Dodge IA) and 4 mg/kg xylazine (Mobay Corporation, Shawneee,
KS). Virginia opossums were anaesthetized with an IM injection of tiletamine plus zolazepam (Telazol®, 5 mg/kg body weight, Aveco Co., Fort Dodge, IA). Approximately
1 mL of blood was aseptically collected from the jugular vein of raccoons and 125 μL from the medial saphenous vein of opossums into ethylenediaminetetraacetic acid
(EDTA) tubes at 3, 7, 10, 14, 17, 21, 24, 28, 35, 42, 49, 56, 70, 84, and 112 days post inoculation (DPI). One raccoon from each group was euthanized at 28 DPI and 112 DPI 92 as representative acute and chronic infections, respectively. One opossum from each group was euthanized at 28 DPI, 56 DPI, and 112 DPI as representative acute, late acute, and chronic infections, respectively. Animals were humanely euthanized under anesthesia by intracardiac injection of sodium pentobarbital (1mg/kg; Butler Company, Columbus,
OH) and exsanguination.
Direct and molecular detection of T. cruzi
At each sampling time, parasitemias were determined by examining 5 µL of whole blood under an 18mm cover glass at 400X magnification with a compound microscope. The entire volume of blood was scanned and the number of counted parasites converted to parasites/mL.
DNA was extracted from 100 µl of whole blood using the DNeasy blood and tissue kit (Qiagen, Inc., Valencia, CA) following the manufacturer‘s protocol. Extracted
DNA was used as template in polymerase chain reaction (PCR) amplification of the D7 divergent domain of the 24Sα rDNA gene of T. cruzi in raccoon infections using a modified nested reaction with primers D75 and D76 (Briones et al., 1999) in the primary reaction and primers D71 and D72 in a secondary reaction (Souto et al., 1996). Because this protocol amplified opossum DNA, opossums were tested for the T. cruzi kinetoplast minicircle DNA by using primers S35 and S36 as previously published (Vallejo et al.,
1999). Total volume of each reaction mixture was 25 μL and contained 5X Buffer, 2 μM of each dNTP, 1 μM of each primer, 2.5 mM MgCl2, and 1.25 U of GoTaq Taq polymerase (Promega Corporation, Madison, WI). The temperature and cycling profile was previously described (Vallejo et al., 1999; Souto et al., 1996). Stringent protocols and controls were used in all PCR assays to prevent and to detect contamination. DNA 93 extraction, amplification, and product analysis were performed in separate dedicated laboratory areas. A negative water control was included in each set of extractions and
PCR reactions as contamination controls. The 330-bp minicircle or 125- or 110-bp 24Sα amplicons were visualized on an ethidium bromide stained 1.5% agarose gel by transillumination.
After euthanasia, animals were necropsied and portions of major tissues
[retropharyngeal lymph nodes, diaphragm, heart, lungs, liver, spleen, gastrointestinal tract, pancreas, kidney, adrenal glands, reproductive organs, urinary bladder, quadriceps muscle, bone marrow, brain, and anal sacs (opossums only)] were collected. One portion of each sample was preserved in 10% neutral buffered formalin for histopathologic examination and the remaining portion stored at -20C until PCR analysis. Frozen tissues were thawed and one 25-mg section of each was aseptically excised. DNA was isolated from tissue using the DNeasy blood and tissue kit (Qiagen) following the manufacturer‘s protocol with a 24 hr tissue lysation step, and used as template following the same PCR parameters described above.
Serology
Indirect immunofluorescent antibody assay was performed as previously described (Yabsley et al., 2001) with plasma at a 1:40 dilution. Briefly, epimastigotes were fixed to serology slides (Erie Scientific, Portsmouth, NH, USA) by air drying and fixation in an acetone wash for 10 minutes. Diluted serum samples and positive and negative controls were added to respective wells and incubated for approximately 25 min.
Two five-minute washes with 1 X phosphate buffered solution (PBS) and a five minute distilled water wash were performed and the slides dried. A commercial fluorescin- 94 labeled anti-species IgG antibody (1:50) was added to slides and incubated for approximately 25 minutes. Two five-minute PBS washes were performed and slides counterstained using a final wash of 1.65% Eriochrome Black T (Sigma, St. Louis,
Missouri, USA) in distilled water. Secondary antibody used during raccoon serology was a goat anti-raccoon IgG (Kirkegaard and Perry Laboratories (KPL), Gaithersburg,
Maryland, USA). After the first incubation, opossum samples were incubated with a rabbit anti-opossum IgG (Bethyl Laboratories, Montgomery, Texas, USA), and then a
FITC-labeled anti-rabbit IgG (KPL). A sample was positive for T. cruzi antibodies if the epimastigotes appeared green under fluorescent microscopy, or low positive if red with a green outline. Negative samples appeared red.
Hemoculture
At euthanasia, hemoculture in DH82 macrophages (Yabsley et al., 2004; Hall et al., 2007) was carried out with 1 mL of EDTA-anti-coagulated whole blood and checked daily for the presence of trypomastigotes. Briefly, in a 50 mL tube, approximately 35 mL of ammonium chloride potassium (ACK) lysing buffer was added to blood, gently inverted for 5 minutes, and centrifuged at 1620 x g for 10 minutes. The supernatant was discarded and the procedure repeated. The buffy coat pellet was resuspended in 5-mL of
MEM and added to a confluent monolayer of DH82 cells.
Histopathology
Formalin-fixed tissues were routinely processed, embedded in paraffin, sectioned at 5 μm, and stained with hematoxylin and eosin. Slides were examined by light microscopy and blindly scored. Histologic lesions were scored as mild, moderate, or 95 severe for each tissue. Presence of amastigote nests were also noted in tissues after scanning 40 fields at 400X magnification.
Results
Raccoons
Parasitemias were first detected at 3 DPI in Y (TcIIb) - and dual (TcI and TcIIa) - inoculated raccoons; trypomastigotes were first noted at 7 DPI in TcIIa-inoculated animals and 10DPI in TcI-inoculated animals. Highest parasite counts were observed in dual-inoculated raccoons with TcIIa infections representing the second highest parasitemia (Figure 5.1). Parasitemias peaked between 17 and 28 DPI, with the exception of Y strain, which peaked at 10 DPI. No detectable parasitemia was noted after 35 DPI and 56 DPI for Y- and TcI-inoculated animals, respectively. The loss of detectable parasitemia in the TcI-infected raccoons could have occurred between 56 DPI, when trypomastigotes were last observed, and 70 DPI. Hemoculture results corresponded to the parasitemia data with the chronically TcI- and Y strain-infected raccoons being hemoculture negative and all other animals being hemoculture positive at the day of euthanasia (Table 5.1). Raccoon 475, which was inoculated with Y strain, was humanely euthanized on 21 DPI because it developed acute clinical signs, including hind limb paralysis and shallow, labored breathing. No other animals displayed clinical signs during the experiment.
PCR detection of T. cruzi DNA in blood varied from parasitemia results. All inoculated groups were first PCR positive by 7 DPI (Table 5.1). Additionally, all animals remained PCR positive through the remainder of the study. The amplicon sizes also 96 corresponded with the genotype of the inoculum, and, in the case of dual infections, only
TcIIa size bands (125bp) were observed (data not shown). Clones of T. cruzi isolated from these animals during hemoculture verified that only TcIIa was isolated from dual- exposed raccoons. In general, T. cruzi was detected in numerous tissues of infected animals by PCR (Table 5.2). Interestingly, in chronically infected animals in all experimental groups, no T. cruzi DNA was detected in spleen samples, yet other organs were positive at the chronic stage and spleens from all acutely infected animals were positive. The only other tissue with regular non-detection of T. cruzi DNA was in half of the male testis sampled.
Time to seroconversion varied between experimental groups (Table 5.1). The
TcIIa-inoculated animals were seropositive by 3 DPI. TcI-inoculated raccoons seroconverted between 10 and 17 DPI, dual-inoculated raccoons between 7 and 10 DPI, and Y strain-inoculated raccoons between 10 and 14 DPI. Following seroconversion, animals remained seropositive throughout the remainder of the study.
Histopathologic lesions were most commonly observed in cardiac and skeletal muscle, brain, and liver. Inflammation was mild in all tissues except heart, skeletal muscle, and brain. In general, acute infections resulted in inflammation in a greater number of tissues as compared to chronic infections (Table 5.3). Inflammation in all tissues was composed primarily of lymphocytes and plasma cells with fewer neutrophils and occasional eosinophils and macrophages. In heart, lesions tended to be more severe around the atria and auricles than near the apex. In mildly affected animals, lesions were composed of small multifocal aggregates of inflammatory cells between myocardial fibers. In more severely affected animals, multifocal to coalescing aggregates of 97 inflammatory cells separated and replaced myocardial fibers and were also present in the endocardium and epicardium. Individual myocardial fibers were necrotic as evidenced by swelling, hypereosinophilia, and fragmentation with loss of cross striations. In brain, lesions consisted primarily of glial nodules with perivascular cuffing less commonly observed. Lesions in skeletal muscle resembled those in the heart, although they tended to be less severe in skeletal muscle. Lesions in liver were mild and consisted primarily of periportal aggregates of inflammatory cells. Amastigote nests were identified in all chronically infected animals and the acutely dual-inoculated raccoon (RAC 469).
Amastigotes were most commonly seen in cardiac and skeletal muscle and less commonly in the adrenal gland and pancreas.
Virginia opossums
Parasitemias were only detected in opossums infected with TcI or dual- inoculated. The highest parasitemia occurred in the dual-inoculated group, with the TcI infection being the next highest; however, trypomastigotes were detected first in TcI opossums at 3 DPI compared to dual-inoculated opossums at 7DPI (Figure 5.2). At the final bleed date, TcI infected animals were still parasitemic, while dual-inoculated opossums no longer were parasitemic. Hemoculture confirmed all the above findings; cultures were negative for the TcIIa- or Y strain (TcIIb)-inoculated groups and for the chronic, dually-inoculated opossums.
Attempts to PCR amplify T. cruzi DNA from whole blood revealed that all TcI- and dual-inoculated opossums were PCR positive at 3 DPI and every bleed date thereafter (Table 5.4). T. cruzi DNA was never detected by PCR in any of the TcIIa- inoculated opossums. Interestingly, Y strain (TcIIb)- inoculated animals were PCR 98 positive at 3 DPI and 7 DPI but became PCR negative by 14 DPI and remained negative through the remainder of the study. Amplification of T. cruzi DNA from tissues collected at necropsy (Table 5.5) revealed numerous organs were positive in opossums in the TcI or dual-inoculated groups while all organs from the TcIIa- and Y-inoculated animals were PCR negative for T. cruzi DNA. Chronically infected animals with patent infections throughout the experiment did not have T. cruzi DNA in the lungs or anal sac (Table 5.5).
Serology revealed interesting results; at least two animals in each experimental group seroconverted, including opossums in the TcIIa- and Y strain-inoculated groups
(Table 5.4). Seroconversion of animals in the dual infection group was slightly earlier (10
DPI) than those in the TcI group (14 DPI). Seroconversion was greatly delayed for the
TcIIa group; the opossum euthanized on DPI 28 failed to seroconvert while the other two opossums seroconverted by 35 and 112 DPI. The Y strain group was seroconverted as early as 10 DPI.
Y strain and TcIIa-inoculated opossums had milder histologic lesions than did opossums in the TcI or dual-inoculated groups, but distribution and character of lesions was similar among all groups (Table 5.6). Heart, skeletal muscle, and anal sacs were most commonly affected. In heart, lesions consisted of multifocal to coalescing areas of inflammation that affected the epicardium, myocardium, and endocardium. Inflammation was often more prominent around the atria and auricles than toward the apex of the heart.
Inflammation was composed of a mixture of lymphocytes, plasma cells, neutrophils, and macrophages with fewer eosinophils. In skeletal muscle, there were multifocal aggregates of lymphocytes, plasma cells, and macrophages with fewer neutrophils and rare eosinophils between and surrounding myofibers. Rare individual myofibers were 99 swollen, hypereosinophilic, and fragmented with loss of cross striations (necrotic). In anal sacs, the lumens often contained sloughed epithelial cells and numerous, often necrotic, neutrophils. Variable numbers of neutrophils, lymphocytes, plasma cells, and eosinophils were noted in the submucosa and muscularis. Brain was affected in only three animals but the most common lesion was glial nodules with perivascular cuffs of lymphocytes and plasma cells observed in one animal. Adrenalitis was observed in four animals. However, a similar finding of suppurative inflammation with multinucleated cells was also observed in the adrenal gland of the negative control, so the significance of this is uncertain. Changes in other tissues in the experimentally infected animals were mild and consisted of primarily lymphoplasmacytic periportal inflammation in liver, lymphoplasmacytic and neutrophilic inflammation in the submucosa and tunica muscularis of the urinary bladder, and mild lymphoplasmacytic interstitial and peripelvic inflammation of the kidney. Lesions in the lungs were considered incidental and included mild pulmonary edema (also noted in the negative control) and lipid pneumonia.
Discussion
The biological characteristics of T. cruzi within various hosts are not well understood, especially in wildlife and in association with parasite genotype. Associations between host species and parasite genotype have been previously suggested and are important in understanding both the domestic and sylvatic cycles of T. cruzi (Clark and
Pung, 1994; Briones et al., 1999; Yeo et al., 2005). In the current study this observed relationship was explored through experimental infections of two common wildlife reservoirs from the United States (raccoons and opossums) with distinct genotypes of T. 100 cruzi. These species were chosen because of the high prevalence detected in these two species to other hosts from which T. cruzi has been isolated, including striped skunks
(Mephitis mephitis), gray fox (Urocyon cinereoargenteus), woodrats (Neotoma spp.), and nine-banded armadillos (Dasypus novemcinctus) (Packchanian, 1942; Mckeever et al.,
1958; Ryan et al., 1985; Yaeger, 1988; Barr et al., 1991a; Brown et al., in press; Kjos et al, unpublished). Also, the large number of isolates that have been characterized from naturally-infected raccoons and Virginia opossums suggested an association that needed to be studied experimentally (Clark and Pung, 1994; Barnabé et al., 2001; Roellig et al.,
2008). Differences in susceptibility to distinct genotypes were demonstrated between and within raccoons and Virginia opossums.
For the Virginia opossums, only those animals that were inoculated with the TcI isolate (either solely or dually with TcIIa) developed parasitemias. Even though the opossums inoculated with the Y-strain (TcIIb) were PCR-positive for at least a week after inoculation, no observable parasitemia was noted in direct blood counts. No evidence of infection (PCR, parasite counts, culture, histopathology) was observed in the TcIIa- inoculated opossums; however, two of the three opossums seroconverted after several weeks to months. These differences between the experimental groups may represent differences in the Virginia opossum‘s reservoir competency which supports field-based molecular typing studies (Clark and Pung, 1994; Briones et al., 1999; Barnabé et al.,
2001; Yeo et al., 2005; Roellig et al., 2008). Because only a few isolates were included in the current study, further tests are needed with additional isolates to determine if Virginia opossums can become infected with other raccoon and other TcII isolates. Our data are also consistent with the analysis of other Didelphis spp. from Central and South America 101 where a significant association between genotype and host was revealed (Yeo et al.,
2005; O‘Connor et al., 2007). Experimental infections of South American Didelphis spp. with a Y strain isolate yielded similar results to the current study where no evidence of persistent parasitemia was found and decreased humoral immune responses were noted
(Jansen et al., 1991). Together these experimental and field-based data suggest that
Didelphis spp. are more highly adapted to TcI strains.
Alternatively, our data suggest raccoons are better hosts for TcIIa. While raccoons were able to develop a patent infection when inoculated with TcI and TcII, the highest parasitemia from a single isolate inoculation was seen with TcIIa. In molecular studies of field isolates, both T. cruzi genotypes have been isolated from raccoons (Clark and Pung,
1994; Briones et al., 1999; Yeo et al., 2005; Roellig et al., 2008), but of the 79 field isolates analyzed in these previous studies, only three were characterized as TcI. Our experimental data supported these field findings.
Possible reasons for the difference in infectivity of certain genotypes to specific hosts may be indicative of biological differences between genotypes, differences in host susceptibility to infection, and an interaction between both isolate characteristics and host susceptibility. Previous in vivo biological characterization studies have reported discordant results with some TcI isolates causing higher parasitemias or infectivity in murine models than TcII (Bértoli et al., 2006; Sanchez et al., 1990); another reports TcI resulting in lower parasitemias than TcII (Lisboa et al., 2007). An in vitro study comparing biological characteristics of Virginia opossum, armadillo, and domestic dog T. cruzi isolates from Louisiana demonstrated differences in cell adhesion and interiorization of trypomastigotes (Barr et al., 1990). The opossums and armadillo 102 isolates had similar protein profiles representing TcI but patterns different from the dog isolate, suggesting different zymodemes or genotypes. Experimental infections in dogs with the same isolates demonstrated dogs develop clinical disease in response to opossum and armadillo isolate inoculation, but not from the dog isolate (Barr et al., 1991b), suggesting differences in infectivity to different host species and biological characteristics. These findings correlate to this study in that the biological characteristics alone may have resulted in different infection dynamics observed between experimental groups, and thus genotypes, in a species. However, the most likely explanation for the differences detected between raccoons and Virginia opossums and between experimental groups is an interaction between the isolate of a particular genotype and a specific host‘s response or innate susceptibility.
Contrasting previous histopathologic findings in wild animals (Barr et al., 1991;
Pietrzak and Pung, 1998), individuals in our study had more severe lesions, particularly during the acute stage of infection. In wild raccoons, mild multifocal and interstitial inflammation was observed in the heart (Pietrzak and Pung, 1998), whereas mild to severe inflammation was observed in cardiac muscle of our experimental raccoons.
Similar findings have been demonstrated in wild-trapped Virginia opossums where no lesions other than mild inflammation were observed (Barr et al., 1991). Wild animals from both of these surveillance studies were likely in the chronic stage of infection. The limited lesions observed is consistent with our data, where less severe lesions were observed in animals sampled at 112 DPI compared with 28 and 56 DPI. Supporting our severe lesions during the acute stage, an acute American trypanosomiasis case was reported in a striped skunk (Mephitis mephitis) with moderate meningitis and subacute to 103 chronic multifocal myocarditis (Ryan et al., 1985). Similar lesions were found in our acute stage experimental animals.
An interesting phenomenon observed was the development of higher parasitemias in opossums and raccoons that were inoculated with two strains of the parasite, even though one strain failed to produce infections in singly-inoculated animals suggesting some type of interaction between parasite strains affected the outcome of infection.
Intraspecific interactions may be responsible for the high parasitemia for both dual- infected species and earlier seroconversion, in the case of the opossums. Mixed infections of genotypes have been described in vectors (Bosseno et al., 1996), mammalian hosts
(Herrera et al., 2005; Lisboa et al., 2006; Herrera et al., 2008; Roellig et al., 2008), and humans (Lauria-Pires et al., 1996), although detection of multiple strains in a single host is rare. In this study, while simultaneous inoculation occurred, one genotype (TcIIa) was detected in raccoons having TcIIa. Mixed infections may not be maintained when animals are co-infected with two genotypes. However, the inoculation of one genotype seemed to alter the infection dynamics of the other isolate. Similar results in opossums could not be determined because T. cruzi could not be isolated from the chronic individual for cloning and analysis using the D7 divergent domain of the 24Sα rDNA gene.
The current study provides evidence that a native wildlife reservoir of the United
States can develop an infection with a non-native strain of T. cruzi. Similar to results of
TcIIa inoculations, the Virginia opossums inoculated with TcIIb (Y strain) from South
America failed to develop a detectable parasitemia. However, T. cruzi DNA was amplified during the first week post inoculation. It is unknown if this detected DNA was 104 from circulation of inoculated trypomastigotes or if limited replication occurred. The failure to detect parasite DNA in the TcIIa-inoculated group suggests that parasites are quickly cleared unless they invade cells and undergo replication. In contrast to the
Virginia opossums, raccoons developed a parasitemia when inoculated with the non- native T. cruzi strain (Y, TcIIb); however, the infection dynamics between the two individual raccoons were dramatically different. One raccoon developed a low-level short-term parasitemia (35 DPI) and survived until the end of the study while the other developed a high parasitemia by 10 DPI and subsequently exhibited hind-limb paralysis and difficulty breathing resulting in humane euthanasia on 21 DPI. Histopathology did not reveal any lesions consistent with the severe clinical signs, but inflammation of the diaphragm or myelitis may be responsible. The ability of raccoons to serve as a potential reservoir for non-native strains of T. cruzi has obvious medical and veterinary implications. Previously, a vector (Triatoma protracta) common in the southeastern US has been shown to serve as a competent vector of non-native strains (Theis et al., 1985;
Theis et al., 1987). The presence of a competent vector and reservoir for non-native T. cruzi suggests the possibility for establishment of different strains in the United States.
Acknowledgments
The authors thank Kate McMillan, Mason Savage, Jessica Murdock, and Emily
Brown (SCWDS) for laboratory assistance and the Animal Resources staff at The
University of Georgia College of Veterinary medicine for assistance with raccoon and opossum care. This study was supported by the National Institutes of Health, National
Institute of Allergy and Infectious Disease grant R15 AI067304. 105
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of Chagas Disease: Didelphis species are natural hosts of Trypanosoma cruzi I 110 and armadillos hosts of Trypanosoma cruzi II, including hybrids. International
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111
Table 5.1. Results* of polymerase chain reaction (PCR) amplification of Trypanosoma cruzi 24Sα rDNA D7 divergent domain, indirect immunofluorescence assay (IFA), and hemoculture from experimentally infected raccoons.
Group Sex 3DPI 7DPI 10DPI 14DPI 17DPI 21DPI 24DPI 28DPI 35DPI 42DPI 49DPI 56DPI 70DPI 84DPI 112DPI TcI Rac 461† M -/- +/- +/- +/n.d. +/+ +/n.d. +/n.d. +/+ euth. Rac 462 F -/- +/low+ +/low+ +/n.d. +/low+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+ TcIIa Rac 472† M -/+ +/+ +/+ +/+ +/+ +/n.d. +/n.d. +/+ euth. Rac 471† M -/low+ +/low+ +/+ +/+ +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+ Dual (IIa & I) Rac 469† F -/- +/- +/low+ +/+ +/+ +/n.d. +/n.d. +/+ euth. Rac 468† M -/- +/low+ +/+ +/+ +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+ Y (TcIIb) Rac 475† M -/- +/- +/- +/+ +/+ +/+ euth. Rac 470 M -/- +/- +/low+ +/low+ +/+ +/n.d. +/n.d. n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+
* DPI= days post inoculation, x/x= PCR result/IFA result, + indicates positive, - indicates negative, n.d. = not done, euth.= euthanized on previous bleed date † hemoculture positive and/or parasitemic on day of euthanasia
112
Table 5.2. Polymerase chain reaction (PCR) amplification of Trypanosoma cruzi minicircle gene in tissues collected at necropsy from experimentally infected raccoons. *
Group Sex DPI† LN† Skeletal Heart Lung Liver Spleen GI† Pancreas Adrenal Kidney Bladder Sex Brain BM† muscle Organ TcI Rac 461 M 28 + + + - + + + + + + + - + + Rac 462 F 112 + + + + - - + + - - + + + + TcIIa Rac 472 M 28 + + + + + + + + + + + + + + Rac 471 M 112 n.d. + + + + - + + + + + - + + Dual (IIa & I) Rac 469 F 28 + + + + + + + + + + + + + + Rac 468 M 112 + + + + + - + + + + + + + + Y (TcIIb) Rac 475 M 21 + + + + + + + + + + + - + + Rac 470 M 112 n.d. + + + + - + + + + + + + +
* + indicates positive, - indicates negative, n.d. = not done †DPI= the day post inoculation that animals were euthanized, LN = lymph node, GI = gastrointestinal tract, BM = bone marrow
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Table 5.3. Inflammation scores * of tissues collected at necropsy from experimentally infected raccoons.
Group Sex DPI † LN† Skeletal Heart Lung Liver Spleen GI† Pancreas Adrenal Kidney Bladder Sex Brain muscle Organ TcI Rac 461 M 28 — — S ‡ M M — — — M ‡ M — — M Rac 462 F 112 — — M — M — — — — — — M — TcIIa Rac 472 M 28 n.d. M ‡ Mo ‡ M M — M M ‡ M ‡ — M M M Rac 471 M 112 n.d. — M — M — — — — — — — M Dual (IIa & I) Rac 469 F 28 — M ‡ S ‡ M M — — — — — — n.d. M Rac 468 M 112 — — Mo ‡ M M — — M — — M — M Y (TcIIb) Rac 475 M 21 — Mo ‡ S — M — M — M — n.d. — Mo Rac 470 M 112 n.d. M — M M — — — — — — — — Neg. Control Rac 473 M 42 — — — — — — — — — — — — —
* n.d.= not determined, — =None observed, M= mild, Mo= moderate, S= severe † DPI, the day post inoculation that animals were euthanized, LN= Lymph node, GI= gastrointestinal tract ‡ Amastigote nest(s) identified in tissue.
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Table 5.4. Results* of polymerase chain reaction amplification (PCR) of Trypanosoma cruzi minicircle, indirect immunofluorescence assay (IFA), and hemoculture from experimentally infected Virginia opossums.
Group Sex 3DPI 7DPI 10DPI 14DPI 17DPI 21DPI 24DPI 28DPI 35DPI 42DPI 49DPI 56DPI 70DPI 84DPI 112DPI TcI Opo 7549† F n.d. +/- +/- +/low+ +/+ +/n.d. +/n.d. +/+ euth. Opo 7539† M +/- +/- +/- +/+ +/+ +/n.d +/n.d +/n.d +/n.d +/n.d +/n.d +/+ euth. Opo 7538† M +/- +/- +/- +/+ +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d +/+ TcIIa Opo 7535 M -/- -/- -/- -/- -/- -/- -/- -/- euth. Opo 7547 F -/- -/- -/- -/- -/- -/- -/- -/- -/low+ -/low+ -/low+ -/low+ euth. Opo 7541 F -/- -/- -/- -/- -/- -/- -/- -/- -/- -/- -/- -/- -/- -/- -/low+ Dual (IIa & I) Opo 7544† M n.d. +/- +/low+ +/low+ +/+ +/n.d. +/n.d. +/+ euth. Opo 7546† F +/- +/- +/+ +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+ euth. Opo 7545 M +/- +/- +/- +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. -/+ Y (TcIIb) Opo 6797 F +/- +/- -/- -/low+ -/+ -/n.d. -/n.d. -/+ euth. Opo 7536 F +/- +/- -/+ -/+ -/+ -/n.d. -/n.d. -/n.d. -/n.d. -/n.d. -/n.d. -/+ euth. Opo 7537 F +/- +/- -/+ -/+ -/+ -/n.d. -/n.d. -/n.d. -/n.d -/n.d. -/n.d -/n.d. -/n.d. -/n.d. -/+
* DPI= days post inoculation, x/x= PCR result/IFA result; + indicates positive, - indicates negative, n.d. = not done, euth.= euthanized on previous bleed date † hemoculture positive and/or parasitemic on day of euthanasia
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Table 5.5. Polymerase chain reaction (PCR) amplification of Trypanosoma cruzi minicircle gene in tissues collected at necropsy from experimentally infected Virginia opossums. *
Group Sex DPI LN† Skeletal Heart Lung Liver Spleen GI† Pancreas Adrenal Kidney Bladder Sex Brain Anal BM† † muscle gland Organ Sac TcI Opo 7549 F 28 + + + - + + - + + - + + - + + Opo 7539 M 56 + + + + + + + + + + + + + + + Opo 7538 M 112 + + + - + + + + + + + + + - + TcIIa Opo 7535 M 28 ------Opo 7547 F 56 ------Opo 7541 F 112 ------Dual (IIa & I) Opo 7544 M 28 - + + + + + + + + + + + + + + Opo 7546 F 56 + + + - + + + + + + + - + + + Opo 7545 M 112 + + - - + + + ------Y (TcIIb) Opo 6797 F 28 ------Opo 7536 F 56 ------Opo 7537 F 112 ------
* + indicates positive, - indicates negative, n.d. = not done † DPI, the day post inoculation that animals were euthanized, LN = lymph node, GI = gastrointestinal tract, BM = bone marrow
116
Table 5.6. Inflammation scores* of tissues collected at necropsy from experimentally infected Virginia opossums.
Group Sex DPI† LN† Skeletal Heart Lung Liver Spleen GI† Pancreas Adrenal Kidney Bladder Sex Brain Anal muscle gland Organ Sac TcI Opo 7549 F 28 — Mo Mo M M — — M M — n.d. — — S Opo 7539 M 56 — M M — M — — M — — M — — S Opo 7538 M 112 — M‡ Mo M — — M — — — — — Mo — TcIIa Opo 7535 M 28 — M M M M — — — — M — — — M Opo 7547 F 56 — — M — — — — — — M — — — n.d. Opo 7541 F 112 — — — M — — — — — — — — — — Dual (IIa & I) Opo 7544 M 28 — M S‡ — — — — — — — M — — n.d. Opo 7546 F 56 — Mo Mo — M — — — Mo Mo M — — Mo Opo 7545 M 112 — M Mo — M — n.d. M — — n.d. — — M Y (TcIIb) Opo 6797 F 28 — — M — M — — — M — — — M Mo Opo 7536 F 56 — — Mo — — — — n.d. — — — — M Mo Opo 7537 F 112 — M M M — — — — M — n.d. — — n.d. Neg. Control Opo 7550 F 112 — — — M — — — — Mo — — — — n.d
* n.d.= not determined, — =None observed, M= mild, Mo= moderate, S= severe † DPI= the day post inoculation that animals were euthanized, LN= Lymph node, GI= gastrointestinal tract ‡ Amastigote nest(s) identified in tissue. 117
Figure 5.1. Parasitemias of individual raccoons (Procyon lotor) experimentally inoculated with different genotypes of Trypanosoma cruzi.
118
Figure 5.2. Parasitemias of individual Virginia opossums (Didelphis virginiana) experimentally inoculated with TcI and dual (IIa & I) genotypes of Trypanosoma cruzi.
119
CHAPTER 6
EXPERIMENTAL INFECTIONS OF TWO SOUTH AMERICAN RESERVOIRS
WITH FOUR DISTINCT TRYPANOSOMA CRUZI STRAINS1
1 Roellig, D.M., McMillan K., Ellis, A.E., VandeBerg, J.L., Champagne, D.E., Yabsley, M.J. submitted to Parasitology, 09/15/2009 120
Summary
Trypanosoma cruzi (Tc), causative agent of Chagas disease, is a diverse species with two primary genotypes, TcI and TcII, with TcII further subdivided into five subtypes (IIa-e). This study evaluated infection dynamics of four genetically and geographically diverse T. cruzi strains in two South American reservoirs, degus (Octodon degus) and gray short-tailed opossums (Monodelphis domestica). Based on prior suggestions of a genotype-host association, we hypothesized that degus (placental) would more readily become infected with TcII strains while short-tailed opossums (marsupial) would be a more competent reservoir for a TcI strain. Individuals (n= 3) of each species were intraperitoneally inoculated with T. cruzi trypomastigotes of TcIIa [North America
(NA)-raccoon (Procyon lotor) origin], TcI [NA-Virginia opossum (Didelphis virginiana)], TcIIb [South America (SA)-human], TcIIe (SA-Triatoma infestans), or both
TcI and TcIIa. Parasitemias in experimentally infected degus peaked earlier [7-14 days post inoculation (DPI)] compared with short-tailed opossums (21-84 DPI). Additionally, peak parasitemias were higher in degus; however, duration of detectable parasitemias for all strains, except TcIIa, was greater in short-tailed opossums. Infections established in both host species with all genotypes, except for TcIIa, which did not establish a detectable infection in short-tailed opossums. These results indicate that both South
American reservoirs support infections with these North and South American isolates; however, infection dynamics differed with host and parasite strain.
Keywords: Trypanosoma cruzi, experimental infection, reservoir host, infection dynamics, degu, gray short-tailed opossum, trypanosome 121
Introduction
Trypanosoma cruzi (Tc), the causative agent of American trypanosomiasis
(Chagas‘ disease), is a hemoflagellate protozoan parasite endemic in the Americas. This parasite species has considerable genetic diversity among isolates and the ability to infect a number of mammalian hosts. A significant factor in new T. cruzi infection occurrence and prevalence is related to the nearly 200 animal reservoirs that have been identified in the Americas (Barretto and Ribeiro, 1979). This provides an opportunity for development of great genetic variability and distinct transmission cycles, host or habitat speciation among triatomine vectors, and changes among mammal and marsupial community structures.
The genetic structure of the T. cruzi population is divided into two primary genotypes, TcI or TcII, with type II having five subtypes (a-e). In South America, all six phylogenetic lineages are present, while only TcI and TcIIa have been identified in the
United States (Clark and Pung, 1994; Barnabé et al., 2001; Hall et al., 2007; Roellig et al., 2008). There is increasing evidence that certain reservoir hosts maintain distinct or certain parasite genotypes. In general, it is suggested that marsupial reservoirs more readily harbor TcI, while placental mammals maintain TcII genotypes (Briones et al.,
1999; Yeo et al., 2005; Roellig et al., 2008).
In the current study, infection dynamics of experimental T. cruzi infection were studied in two natural wild reservoirs from South America, degus (Octodon degus) and gray short-tailed opossums (Monodelphis domestica). The degu is a diurnal, highly social caviomorph rodent native to Chile. In previous studies, prevalence in wild-captured degus in different geographic regions ranged from 2.2% to 21.4% by hemagglutination or 122 xenodiagnosis and 51%-61% by molecular detection methods (Whiting, 1946; Duran et al., 1989; Jimenez and Lorca, 1990; Rozas et al., 2005; Rozas et al., 2007). The gray short-tailed opossum is a solitary, terrestrial, didelphid opossum that is native to Bolivia,
Brazil and Paraguay. A characteristic of short-tailed opossums, as well as some other
South American marsupials, is the absence of a pouch for neonates, which may have implications in transmission and maintenance of T. cruzi. The underdeveloped young cling to the teats of females during their development. Prevalence data for wild-captured gray short-tailed opossums range from 11.1% to 66.7% in different geographic regions by various detection methods, such as hemoculture, xenodiagnosis, and serology (Herrera et al., 2005; Roque et al., 2008).
Genotyping of T. cruzi isolated from both species has been previously reported, and multiple genotypes have been identified in both species. Only six isolates have been characterized from short-tailed opossums; two were TcII (undefined subtype), one was
TcIIc and three were TcI (Yeo et al., 2005; Roque et al., 2008). Multiple genotypes have also been reported from naturally infected degus (Yeo et al., 2005; Campos et al., 2007;
Rozas et al., 2007; Spotorno et al., 2008; Galuppo et al., 2009). One study showed that
TcI and TcIIb were more prevalent than other detected genotypes (TcIId and TcIIe)
(Rozas et al., 2007), while another found higher prevalence of TcIId than TcIIb and TcI genotypes (Galuppo et al., 2009). These field-based studies suggest that these two hosts are susceptible to both major genotypes of T. cruzi, which is in contrast to data from two major North American reservoirs which exhibit significant predilections for certain T. cruzi genotypes (Roellig et al., 2008; Roellig et al., 2009). 123
The goals of the present study were to ascertain reservoir potential of M. domestica and O. degus for four different T. cruzi genotypes from the United States and
South America. In addition, the infection dynamics of these four different strains were studied to determine if there are any associations between parasite genotype and host species.
Materials and Methods
The two North American isolates used in this study were originally isolated from a naturally-infected Florida raccoon, Procyon lotor [FL-RAC9 (TcIIa)], and Virginia opossum, Didelphis virginiana [FL-OPO3 (TcI)], from northwestern Florida (Roellig et al., 2008). Two South American strains, Y (TcIIb) and Tulahuen (TcIIe), were generously provided by Drs. Rick Tarleton and Roberto DoCampo (The University of
Georgia, Athens, GA, USA), respectively. Each strain was molecularly typed as previously described (Brisse et al., 2001; Roellig et al., 2008). Epimastigotes were passaged from Liver-Infusion Tryptose (LIT) medium into DH82 canine macrophage monolayer as previously described (Roellig et al., 2009).
Laboratory-raised short-tailed opossums were acquired from the Southwest
Foundation for Biomedical Research (San Antonio, TX) and individually housed in large rodent cages in climate-controlled animal housing at the College of Veterinary Medicine,
University of Georgia (Athens, GA). Captive-bred degus were purchased from a commercial source (S&S exotics, Houston, TX) and similarly housed at a maximum of three individuals per cage. All animals were cared for in accordance with the guidelines 124 of the Institutional Animal Care and Use Committee and under an animal use protocol approved by this committee at the University of Georgia.
For both species, animals were randomly separated into five experimental groups and one negative control group. Individuals in groups 1, 2, 3, or 4 (n=3) were inoculated intraperitoneally (IP) with 1x106 culture-derived trypomastigotes of FL-OPO3 (TcI), FL-
RAC9 (TcIIa), Y (TcIIb), or Tulahuen (TcIIe) strain, respectively. Group 5 individuals
(n=3) were inoculated with 5x105 FL-OPO3 strain and 5x105 FL-RAC9 strain trypomastigotes in single mixed inoculum. Negative controls (n=2) for both species were similarly inoculated with an equivalent volume of MEM.
For handling and blood collection, animals were anaesthetized with subcutaneous administration of a mixture of ketamine (Fort Dodge Laboratories, Inc., Fort Dodge, IA) and xylazine (Mobay Corporation, Shawnee, KS). Degus were given 100mg/kg of ketamine and 20 mg/kg of xylazine; short-tailed opossums were given 50 mg/kg ketamine and 10 mg/kg xylazine. Approximately 20 to 100 µL of blood were aseptically collected at 0, 7, 14, 21, 28, 35, 42, 49, 56, 70, 84, and 112 days post inoculation (DPI) from the medial saphenous vein of degus and lateral tail vein of short-tailed opossums using heparinized capillary tubes. One animal from each group was euthanized at 28, 56, and 112 DPI, representative of acute, late acute, and chronic infections, respectively.
Animals were humanely euthanized under anesthesia by CO2. After euthanasia, animals were exsanguinated via cardiac-puncture and blood was collected into 4 mL vacuette ethylenediaminetetraacetic acid (EDTA) tubes (Greiner Bio-one, Monroe, NC).
At each sampling time, parasitemias were determined by examining 5μL of whole blood as previously described (Roellig et al., 2009). For each time point, DNA was 125 extracted from 20 to 100 μL of red blood cell/buffy coat homogenate using the GFX genomic blood DNA purification kit (Amersham Biosciences, Piscataway, NJ) or
DNeasy blood and tissue kit (Qiagen, Inc., Valencia, CA) following the manufacturers‘ protocols. After euthanasia, animals were necropsied and portions of major tissues
(retropharyngeal lymph nodes, diaphragm, heart, lungs, liver, spleen, gastrointestinal tract, pancreas, kidney, adrenal glands, reproductive organs, urinary bladder, quadriceps muscle, bone marrow, and brain) were collected. DNA was isolated from tissue using the
DNeasy blood and tissue kit (Qiagen) following the manufacturer‘s protocol with a 24 hr tissue digestion step. Extracted DNA was used as template in a modified nested PCR amplification of the T. cruzi 24Sα rDNA D7 divergent domain (Souto et al., 1996) as previously published (Roellig et al., 2009). Negative samples were verified by amplification of the size-variable domain of the 18S rRNA gene (Clark and Pung, 1994) as previously described (Brisse et al., 2001).
Blood (0.5 mL) collected at euthanasia was cultured in DH82 cells (Yabsley et al.,
2004; Hall et al., 2007). Cultures were checked daily for the presence of trypomastigotes.
For xenodiagnosis of chronically-infected animals, laboratory-raised Rhodnius prolixus nymphs (4th and 5th instars) (n=3) were fed until repletion on each anesthetized, chronically infected animal from each group. Bugs were allowed to digest the blood meal and molt in an isolated, temperature and humidity controlled environment. The intestinal tract of resultant 5th instars or adults were removed, added to 700 μL of PBS, vortexed, and boiled for 15 min. This solution was used for PCR amplification of kinetoplast DNA as described above. 126
Indirect immunofluorescent antibody assays were performed as previously described (Yabsley et al., 2001; Roellig et al., 2009) with plasma at a 1:40 dilution.
Secondary antibody used during degu serology was a FITC-labeled goat anti-mouse IgG
(Kirkegaard and Perry Laboratories (KPL), Gaithersburg, Maryland, USA). After the first incubation, short-tailed opossum samples were incubated with a rabbit anti-opossum IgG
(Bethyl Laboratories, Montgomery, Texas, USA), and then a FITC-labeled anti-rabbit
IgG (KPL). A sample was positive for T. cruzi antibodies if epimastigotes appeared green under fluorescent microscopy, or weak positive if red with a green outline. Negative samples appeared red.
Formalin-fixed tissues were routinely processed, embedded in paraffin, sectioned at 5 μm, and stained with hematoxylin and eosin. Slides were examined by light microscopy and blindly scored by a veterinary pathologist. Histologic lesions were scored as mild, moderate, or severe for each tissue. The presence of amastigote nests was also noted in tissues after scanning 40 fields at 400X magnification.
Results
Degus
Parasitemias were first detected in all animals at 7 DPI (Figure 6.1). Highest parasite counts were observed in animals inoculated with TcIIb and TcI, and all animals had a rapid decline in parasitemia. Significant differences in parasitemias during the first
28 days of infection were noted between experimental groups (F=13.65, p<0.001) as determined by Greenhouse-Geiser MANOVA methodology. Parasites were not observed in the TcIIb- and TcIIa-inoculated animals after 28 DPI; parasites were undetectable after 127
56 DPI for the TcI, TcIIe, and dual-infected groups. At 28 DPI, all acutely infected animals still had detectable parasitemia at euthanasia and were also hemoculture positive
(Table 6.1). At 56 DPI, late acute phase animals had variable positive results with TcIIa- and dual-inoculated animals being hemoculture negative, while TcI- and TcIIe-inoculated animals were hemoculture positive. At 112 DPI, none of the chronically infected degus had detectable parasitemias; however, hemoculture and xenodiagnosis indicated that
TcI-, TcIIe-, and dual (TcI/TcIIa)-inoculated animals were still parasitemic.
Molecular detection of T. cruzi DNA in blood samples was achieved for all experimental groups. Animals were PCR positive by 7 or 14 DPI, with the exception of one degu, which was only positive at 35 DPI (Table 6.1). That individual degu also had a low parasitemia at 7 and 14 DPI, and parasites were not observed in blood on other days.
PCR amplification was intermittent for many of the animals with no detection on some bleed days, however, trends could be observed. All experimental groups were PCR positive through the acute phase (28 DPI), but T. cruzi DNA was only amplified in dual- and TcIIe-chronically infected animals (112 DPI). Amplification of T. cruzi DNA in tissue samples was achieved for all animals (data not shown). For many of the animals, all tissues were PCR positive but the hearts, quadriceps, and spleens were PCR positive for all animals. Serology revealed seroconversion of all animals by 14 DPI and all remained seropositive at the time of euthanasia (Table 6.1).
Lesions were common in heart (n=14), skeletal muscle (n=13), brain (n=11), kidney (n=9), and urinary bladder (n=9). Lesions were occasionally noted in liver (n=6), pancreas (n=5), adrenal gland (n=4), testicle (n=4), lung (n=2), and intestine (n=2). In heart and skeletal muscle, lesions consisted of myofiber necrosis and multifocal 128 aggregates of lymphocytes and plasma cells with occasional macrophages or neutrophils.
Inflammation in skeletal muscle was mild (n=10) to moderate (n=3) and ranged from mild (n=5) to moderate (n=5) to severe (n=4) in heart. Lesions in brain were mild (n=7) to moderate (n=4) and included lymphoplasmacytic perivascular cuffing, glial nodules, and meningitis. Pseudocysts or amastigotes were observed in multiple tissues including heart (n=7), skeletal muscle (n=3), brain (n=1), testicle (n=1), intestine (n=1), adrenal gland (n=1), and urinary bladder (n=1).
Short-tailed opossums
Parasitemias were detected in TcIIb-, TcI-,TcIIe-, and dual-infected animals by 7
DPI, but TcIIa-inoculated animals never developed a detectable parasitemia (Figure 6.2).
No significant differences in parasitemias between experimental groups were detected by
Greenhouse-Geiser MANOVA methodology (F=7.2716, p=0.1207). TcIIb-infected animals were parasitemic until 28 DPI, TcI- and dual-infected opossums until 84 DPI, and TcIIe-infected opossums until 112 DPI. Hemoculture and xenodiagnosis confirmed that TcIIa-infected animals were either not parasitemic or had parasitemias below detection limits (Table 6.2). The TcIIb-acutely infected opossum was not parasitemic or hemoculture positive on the day of euthanasia. Interestingly the other two animals in this group were hemoculture positive although parasites could not be found in the 5 μL of blood examined following DPI 28. All other groups had circulating parasitemias that were detected by hemoculture, parasite counts, and/or xenodiagnosis.
Similar to parasitemia determination, hemoculture, and xenodiagnosis, PCR amplification attempts in TcIIa-inoculated animal blood failed to yield positive results
(Table 6.2). The most consistent detection of T. cruzi DNA was accomplished for TcIIe- 129 and TcI-inoculated animals. At least one individual for each of these two groups was first
PCR positive on 7 DPI. TcIIb and dual-inoculated groups were also PCR positive by 7
DPI, however, T. cruzi DNA was only detected intermittently.
PCR amplification in tissues yielded similar results, with tissues from TcIIa- inoculated animals all being negative for T. cruzi by PCR while all other groups had at least one PCR-positive tissue (data not shown). T. cruzi DNA was most commonly amplified from skeletal muscle (quadricep and diaphragm). There were no differences in detection among tissues of animals within an experimental group during different stages of infection. Additionally, no differences in amplification were observed among experimental groups. Serology revealed all animals seroconverted by 21 DPI, with animals in TcIIb- and TcIIa-inoculated groups seroconverting after the TcI group.
Histologic lesions were uncommon and were usually mild. Heart was the only tissue consistently affected with 10 of 16 animals having myocardial lesions. Lesions were observed rarely in brain (n=2), pancreas (n=2), liver (n=2), adrenal gland (n=2), kidney (n=1), intestine (n=1), urinary bladder (n=1), and skeletal muscle (n=1). In all organs, inflammation was primarily lymphoplasmacytic with fewer histiocytes and occasional neutrophils and eosinophils. One opossum had a single glial nodule as the only lesion in the brain. Pseudocysts were not observed in any of the tissues examined.
Discussion
The maintenance and continuation of the T. cruzi sylvatic cycle is dependent on a competent vector feeding on a parasitemic animal. Since T. cruzi is a genetically and biologically diverse species that can infect a wide range of mammalian hosts, it is 130 reasonable to hypothesize that certain animal species may maintain parasitemias longer than others and, thus, have differences in their ability to serve as reservoirs. These differences in reservoir potential may be based on the host species or genetic makeup, the genotype of the parasite, or a combination of both. In this preliminary study, the reservoir potential and infection dynamics of experimental T. cruzi infections in degus and short- tailed opossums were investigated by inoculating animals with different T. cruzi genotypes.
Similar to experimental infections with raccoons, another placental mammal
(Roellig et al., 2009), degus developed patent infections after inoculation with each of the four isolates (representing genotypes TcI, TcIIa, TcIIb, and TcIIe) analyzed in this study.
These data support molecular typing studies conducted on isolates from naturally- infected degus from South America that showed natural infection with multiple genotypes including TcI, TcIIa, TcIIb, TcIId, and TcIIe singly, and some mixed infections (Yeo et al., 2005; Campos et al., 2007; Rozas et al., 2007; Spotorno et al.,
2008). Interestingly, two genotypes, TcI and TcIIe, maintained parasitemias during chronic infections (112 DPI) that were sufficient to infect xenodiagnostically fed R. prolixis. Because of their ability to maintain parasitemias for a long period of time, degus may be considered important reservoirs for the two genotypes (TcI and TcIIe). Further experimental studies with additional strains and larger sample sizes would help to understand the infection dynamics of T. cruzi in degus and identify infectivity and maintenance differences between genotypes suggested in this study.
In the case of short-tailed opossums, animals inoculated with TcIIa seroconverted, but a patent infection could not be detected by any other means including molecular and 131 direct examination of blood and tissues. Findings were similar to experimental and field- based molecular studies that found another marsupial, the Virginia opossum (Didelphis virginiana), do not maintain infections with TcIIa (Clark and Pung, 1994; Barnabé et al.,
2001; Roellig et al., 2008; Roellig et al., 2009). However, field isolates of other marsupial species from South America, including D. marsupialis and P. frenata, indicate that these species can be infected with TcI, TcIIa, and TcIIc South American genotypes based on zymodeme analysis (Miles et al., 1981; Póvoa et al., 1984; Pinho et al., 2000).
Additionally, M. brevicaudata has been shown to be naturally infected with South
American TcIIa and TcIIc genotypes (Miles et al., 1981; Póvoa et al., 1984). Differences in infectivity of TcIIa strains from South versus North America in marsupials may be indicative of biological differences in the parasite and not host susceptibility.
Our findings suggest that short-tailed opossums may serve as reservoirs for multiple T. cruzi strains, including TcI, TcIIb, TcIIe, and mixed infections. These data expand our knowledge on the genotypes to which gray short-tailed opossums are susceptible, which were previously limited to TcIIc and TcI based on natural infections reported from the Gran Chaco of Paraguay and Redencão, Brazil (Yeo et al., 2005;
Roque et al., 2008). The peak in parasitemia seen in the TcI-inoculated animal at 84 DPI is believed to be an artifact of differences between experimental animals in this group as one of two animals was euthanized at the previous time point. As all chronically-infected experimental animals were parasitemic at the time of euthanasia and no differences were statistically detected during acute infection, short-tailed opossums appear to develop long-term parasitemias with multiple genotypes, which is in contrast to Virginia opossums that were inoculated with multiple strains (Roellig et al., 2009). 132
The current study also demonstrated that degus and short-tailed opossums are competent hosts for North America T. cruzi strains. Similar to findings that North
American hosts can become infected with South American isolates, no differences in infectivity based on the geographical origin of the isolates were observed (Roellig et al.,
2009). Field studies have often found that both major lineages (TcI and TcII) can infect vector species (Marcet et al., 2005; Falla et al., 2009), and experimental studies, including the current one, have described infection in vectors susceptible to multiple genotypes (Perlowagora-Szumlewicz et al., 1990; Coronado et al., 2006; Campos et al.,
2007). Combined with the experimental data from the present study, there is a potential for a non-native strain to become established in South America.
This study suggests different T. cruzi genotypes induce distinct infection dynamics in divergent host species. Further work with additional isolates and genotypes and greater sample sizes will enable a better understanding of parasite genotype-host interactions. Such information would be vital for understanding the epidemiology and epizootiology of Chagas‘ disease and may lead to better preventative measures in endemic regions.
Acknowledgments
The authors thank Mason Savage, Jessica Murdock, Wendy Fujita, and Emily
Brown (SCWDS) for laboratory assistance and the Animal Resource staff at The
University of Georgia College of Veterinary Medicine for assistance with degu and opossum care.
133
Financial support
This study was primarily supported by the National Institutes of Health, National Institute of Allergy and Infectious Disease Grant R15 AI067304. KM was a Georgia Veterinary
Scholar and part of her support was obtained from T35 RR022685-01A1 from the
National Center for Research Resources (NCRR), a component of the National Institutes of Health (NIH). Its contents are solely the responsibility of the authors and do not necessarily represent the official view of NCRR or NIH. In addition, we thank Merck-
Merial Ltd. and the Veterinary Medical Experimental Station for their financial support of the students and the Georgia Veterinary Scholars program as a whole. Additional support was through funding provided to John L. VandeBerg by the Robert J. Kleberg,
Jr., and Helen C. Kleberg Foundation, and to SCWDS by the Federal Aid to Wildlife
Restoration Act (50 Stat. 917) and through sponsorship of the fish and wildlife agencies of Alabama, Arkansas, Florida, Georgia, Kansas, Kentucky, Louisiana, Maryland,
Mississippi, Missouri, North Carolina, Oklahoma, Puerto Rico, South Carolina,
Tennessee, Virginia, and West Virginia.
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Table 6.1. Results* of polymerase chain reaction (PCR) amplification of Trypanosoma cruzi 24Sα rDNA D7 divergent domain and indirect immunofluorescence assay (IFA) from experimentally infected degus. Group 7 DPI 14 DPI 21 DPI 28 DPI 35 DPI 42 DPI 49 DPI 56 DPI 84 DPI 112 DPI TcIIb 6752-R +/- +/+ +/+ +/+ euth. euth. euth. euth. euth. euth. 6756-2R† +/- +/+ +/+ euth. euth. euth. euth. euth. euth. euth. 6752-L +/- -/weak+ -/+ -/n.d. -/n.d. +/n.d. +/n.d. +/n.d. -/n.d. -/+ TcI 6743-R‡ +/- +/+ +/+ -/n.d. +/n.d. +/n.d. +/n.d. -/n.d. -/n.d. -/+ 6743-L† +/- +/+ +/+ +/n.d. -/n.d. +/n.d. -/n.d. +/+ euth. euth. 6743-B† +/- +/+ +/+ +/+ euth. euth. euth. euth. euth. euth. TcIIa 6749-L +/- +/+ -/+ +/n.d. +/n.d. +/n.d. -/n.d. -/+ euth. euth. 6749-R -/- -/+ -/+ -/n.d. -/n.d. +/n.d. -/n.d. -/n.d. -/n.d. -/+ 6749-B† +/- +/+ -/+ +/+ euth. euth. euth. euth. euth. euth. TcIIe 6754-L†,‡ -/- +/+ +/+ -/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+ 6756-2L† +/- +/+ +/+ +/n.d. +/n.d. -/n.d. -/n.d. -/+ euth. euth. 6754-R† -/- -/weak+ +/+ +/+ euth. euth. euth. euth. euth. euth. Dual (TcIIa & TcI) 6747-L† +/- +/+ +/+ +/+ euth. euth. euth. euth. euth. euth. 6747-R +/- +/+ +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/+ euth. euth. 6747-B‡ +/- +/+ +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. -/+ *DPI=days post inoculation, x/x=PCR result/IFA result, + indicates positive, - indicates negative, n.d. = not done, euth.= euthanized on previous date † hemoculture positive and/or parasitemic on day of euthanasia ‡ positive by xenodiagnosis (determined only for chronic animals) 140
Table 6.2. Results* of polymerase chain reaction amplification (PCR) of Trypanosoma cruzi 24Sα rDNA D7 divergent domain and indirect immunofluorescence assay (IFA) of experimentally infected short-tailed opossums. Group 7 DPI 14 DPI 21 DPI 28 DPI 35 DPI 42 DPI 49 DPI 56 DPI 84 DPI 112 DPI TcIIb 6886 +/n.d. -/- -/weak+ -/+ euth. euth. euth. euth. euth. euth. 6887† +/- -/+ -/+ -/n.d. +/n.d. +/n.d. -/n.d. -/n.d. -/n.d. -/+ 6888† +/- +/+ -/+ -/n.d. +/n.d. +/n.d. +/n.d. -/+ euth. euth. TcI 6889† -/- +/+ -/+ +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+ 6890† +/weak+ +/weak+ +/+ -/n.d. -/n.d. +/n.d. +/n.d. +/+ euth. euth. 6891† +/weak+ +/weak+ +/weak+ +/+ euth. euth. euth. euth. euth. euth. TcIIa 6892 -/- -/weak+ -/weak+ -/n.d. -/n.d. -/n.d. -n.d. -/+ euth. euth. 6893 -/- -/+ -/+ -/n.d. -/n.d. -/n.d. -/n.d. -/n.d. -/n.d. -/+ 6894 -/- -/- -/+ -/+ euth. euth. euth. euth. euth. euth. TcIIe 6898† +/- +/+ -/+ +/+ euth. euth. euth. euth. euth. euth. 6883† -/+ -/+ -/n.d. +/n.d. +/n.d. +/n.d. -/n.d. +/+ euth. euth. 6899† -/- +/+ +/n.d. +/n.d. -/n.d. +/n.d. +/n.d. -/n.d. +/n.d. +/+ Dual (TcIIa & TcI) 6895†, ‡ +/- +/+ +/+ +/n.d. -/n.d. +/n.d. +/n.d. +/n.d. +/n.d. +/+ 6896† +/+ +/+ +/+ +/n.d. +/n.d. +/n.d. +/n.d. +/+ euth. euth. 6897† -/- -/+ +/+ +/+ euth. euth. euth. euth. euth. euth. *DPI=days post inoculation, x/x=PCR result/IFA result, + indicates positive, - indicates negative, n.d. = not done, euth.= euthanized on previous date † hemoculture positive and/or parasitemic on day of euthanasia ‡ positive by xenodiagnosis (determined only for chronic animals) 141
Figure 6.1. Parasitemias of degus (Octodon degus) experimentally inoculated with different genotypes of Trypanosoma cruzi. Black arrows indicate days when an individual from each experimental group was euthanized. Statistical differences in parasitemias were detected between experimental groups through the acute stage of infection (MANOVA, F=13.65, p<0.0001). 142
Figure 6.2. Parasitemias of short-tailed opossums (Monodelphis domestica) experimentally inoculated with different genotypes of Trypanosoma cruzi. Black arrows indicate days when an individual from each experimental group was euthanized. No statistical differences in parasitemias were detected between experimental groups through the acute stage of infection (MANOVA, F=1.9335, p=0.1207)
143
CHAPTER 7
ORAL TRANSMISSION OF TRYPANOSOMA CRUZI WITH OPPOSING EVIDENCE
FOR THE THEORY OF CARNIVORY1
1 Roellig, D.M., Ellis, A.E., Yabsley, M.J. 2009. Journal of Parasitology. 95: 360-364. Reprinted here with permission of publisher. 144
Abstract
We present the first demonstration of oral transmission of Trypanosoma cruzi to raccoons, a natural reservoir host in the United States, by ingestion of trypomastigotes and infected bugs, but not infected tissue. To investigate an alternative, non-vector based transmission method, we tested the hypothesis that raccoons scavenging on infected hosts results in patent infection. Macerated tissue from selected organs infected with amastigote stages of T. cruzi was orally administered to experimental groups of raccoons
(n=2/group) at 2, 12, or 24 hr after collection of the tissue samples. Additionally, raccoons (n=1) in control groups were inoculated intravenously or per os with trypomastigotes. To further elucidate transmission routes of T. cruzi to raccoons, infected
Rhodnius prolixus were fed to raccoons (n=2). Raccoons did not become infected after ingestion of amastigote-infected tissues as evidenced by negative PCR results from blood and tissue, lack of seroconversion, and negative parasitemias. However, per os transmission can occur by ingestion of the infective trypomastigote stage or infected reduviid bugs. We conclude from these findings that oral transmission of T. cruzi may be a route of infection for wildlife in sylvatic cycles, but the scavenging behavior of animals is not likely a significant transmission route.
Trypanosoma cruzi, a hemoflagellate protozoan parasite, has a complex life cycle that includes both domiciliary and sylvatic cycles. Reservoir hosts for the etiologic agent of Chagas disease include a range of wild mammals in endemic regions of the Americas.
In the United States, there have been few documented human cases, but the prevalence of
T. cruzi in US wildlife based on serology, culture isolation, and/or PCR can be equally as high as in South America (Barr et al, 1991). Studies conducted in the southeastern United 145
States indicate that raccoons and opossums have the highest prevalence of T. cruzi compared with other mammals.
Classic transmission of infective trypomastigotes from the vector (triatomine bugs) to mammals requires invasion of the oral, nasal, or ocular mucosa, or invasion through an abrasion or cut in the dermis near the bug defecation site. Many have proposed alternative modes of transmission of T. cruzi in wildlife species because of inconsistent use of dens by animals and the apparent low density of vectors in dens
(Jansen et al, 1994). Experimental per os infection trials in Virginia opossums (Yeager,
1971) and striped skunks (Davis et al., 1980) have implied direct oral transmission as the presumptive natural route with the ingestion of infected triatomid bugs or oral lavage with infected intestinal contents, respectively. Conversely, microcosm experiments have demonstrated that opossums rarely, if ever, prey on infected bugs in simulated dens but nonetheless acquire T. cruzi (Schweigmann et al, 1995).
In addition to the ingestion of vectors, numerous claims have been made about the importance of carnivory in maintenance of the sylvatic cycle (see Dias, 2006; Miles,
2004; Coura 2006) However, no experimental data in natural reservoirs have been produced to support or refute such declarations. The objective of the current study was to elucidate the natural transmission for wildlife reservoir hosts and determine the role of carnivory in T. cruzi transmission by simulating natural eating habits of raccoons. We hypothesized that the ingestion of amastigote-infected tissues would produce a detectable
T. cruzi infection, which would be identified by seroconversion, development of a patent parasitemia, histologic detection of amastigotes, and amplification of T. cruzi DNA from blood and tissues. Additionally, raccoons were expected to develop patent infections after 146 ingestion of infected triatomine bugs and food contaminated with culture-derived trypomastigotes.
Materials and Methods
Inoculation material
T. cruzi isolated from a raccoon trapped in Torreya State Park, FL in 2005 (FL
RAC 9) was used as the inoculation source in the experiments. This isolate was previously shown to be a type IIa, which is the group most often associated with raccoon infections (Roellig et al, 2008). The original isolate was stored in liquid nitrogen prior to culture in Liver-Infusion Tryptose (LIT) medium at the commencement of this study.
Epimastigotes were passaged from Liver-Infusion Tryptose (LIT) medium into DH82 canine macrophage monolayers at 1:5 dilutions to yield the culture-derived- trypomastigotes. Trypomastigotes were pelleted from culture by centrifugation at 1,620 g for 15 min and resuspended in minimum essential medium (MEM). Amastigote- containing tissue was obtained from parasitemic, juvenile raccoons (n=2) 18 days after intravenous (IV) inoculation with 1 x 106 culture-derived trypomastigotes. Heart, spleen, quadriceps muscle, diaphragm, urinary bladder, and liver were collected at necropsy, pooled, and coarsely ground using a tissue grinder. Trypanosoma cruzi infection of tissues was confirmed by polymerase chain reaction (PCR) of individual and ground tissue and observation of pseudocysts in hematoxylin and eosin stained sections of heart tissue (data not shown). Laboratory-reared Rhodnius prolixus nymphs (4th and 5th instars)
(n=6) were fed until repletion on T. cruzi-infected raccoons (n=2) with detectable parasitemias and allowed to digest the blood meal for 19 days. 147
Animals and experimental design
Eleven juvenile raccoons obtained from Ruby Fur Farm, Inc. (New Sharon, Iowa) were housed individually or in pairs in climate-controlled animal housing at the College of Veterinary Medicine, University of Georgia (Athens, Georgia); they were given food and water ad libitum, except food was withheld for 24 hr pre-exposure. All animals used in this study were maintained in accordance with the guidelines of the Institutional
Animal Care and use Committee and under animal use protocol approved by the
Institutional Animal Care and Use Committee at the University of Georgia. Before use, all raccoons were determined to be negative for antibodies reactive with T. cruzi (as described below).
Animals were separated into 6 experimental groups. Group 1 (n=1) served as a positive control for parasite infectivity and was inoculated intravenously (IV) with 1 x
106 culture-derived trypomastigotes. Individuals in group 2, 3, or 4 (n=2) ingested approximately 33.3 g of pooled amastigote-infected tissue that was held at room temperature for 2, 12, or 24 hr post-mortem, respectively. Group 5 raccoons (n=2) each ingested 3 R. prolixus nymphs shedding metacyclics in feces as determined by light microscopy. Group 6 (n=1) was inoculated per os (PO) by feeding 1 x 106 culture-derived trypomastigotes mixed with commercial canned cat food. Two negative control raccoons were inoculated IV with equivalent volumes of media as group 1.
All animals were anesthetized with an intramuscular injection of a mixture of 20 mg/kg ketamine (Fort Dodge Laboratories, Inc, Fort Dodge Iowa) and 4 mg/kg xylazine
(Mobay Corporation, Shawnee, Kansas) for handling and blood collection. Blood samples were aseptically collected from the jugular vein into ethylenediaminetetraacetic 148 acid (EDTA) tubes every 7 days post-inoculation (DPI) until being killed. The raccoon in group 1 was killed at 28 DPI, group 5 at 35 DPI, and all others at 42 DPI. Animals were humanely killed under anesthesia by exsanguination and intracardiac injection of sodium pentobarbital (1mg/kg; Butler Company, Columbus, Ohio).
Direct and molecular detection of T. cruzi
Parasitemias were determined by examining 5 µl of whole blood under an 18-mm cover glass at 400x magnification with a compound microscope. The entire volume of blood was scanned and the number of counted parasites converted to parasites/ml.
DNA was extracted from 100 µl of whole blood using the DNeasy blood and tissue kit (Qiagen, Inc., Valencia, California) following the manufacturer‘s protocol.
Extracted DNA was used as template in PCR amplification of D7 divergent domain of the 24Sα rDNA gene using a modified nested reaction with primers D75 and D76
(Briones et al., 1999) in the primary reaction and primers D71 and D72 in a secondary reaction (Souto et al., 1996). Total volume of each reaction mixture was 25 ul and contained 5x Buffer, 2 uM of each dNTP, 1 uM of each primer, 2.5 mM MgCl2, and 1.25
U of GoTaq Taq polymerase (Promega Corporation, Madison, Wisconsin). The temperature and cycling profile was previously described (Souto et al., 1996). Stringent protocols and controls were used in all PCR assays to prevent and to detect contamination. DNA extraction, amplification, and product analysis were performed in separate dedicated laboratory areas. A negative water control was included in each set of extractions and PCR reactions as contamination controls. The 120bp amplicons were visualized on an ethidium bromide stained 1.5% agarose gel by transillumination. 149
After death, raccoons were necropsied and representative samples of major organs
(retropharyngeal lymph nodes, skeletal muscle (diaphragm and quadriceps), heart, lungs, liver, spleen, gastrointestinal tract, pancreas, kidney, adrenal glands, reproductive organs, urinary bladder, quadriceps muscle, and brain) were collected. One portion of each sample was preserved in 10% neutral buffered formalin for histologic examination and the remaining portion stored at –20 C until PCR analysis. Frozen tissues were thawed and
1, 25-mg section of each was aseptically excised. DNA was isolated from tissue using the
DNeasy blood and tissue kit (Qiagen) following the manufacturer‘s protocol with a 24 hr tissue lysation step.
Serology
Indirect immunofluorescent antibody assay was performed as previously described (Yabsley et al., 2001) with plasma at a 1:40 dilution. Briefly, epimastigotes were fixed to serology slides (Fisher Scientific, Rome, Georgia) by air drying and fixation in an acetone wash for 2 min. Diluted serum samples and positive and negative controls were added to respective wells and incubated for approximately 25 min. Two, 5 min washes with 1x PBS and a 5 min distilled water wash were performed and the slides dried. Diluted FITC-labeled goat anti-raccoon antibodies (Kirkegaard and Perry
Laboratories (KPL), Gaithersburg, Maryland; 1:50) were added to slides and incubated for approximately 25 min. Two, 5 min PBS washes were performed and were counterstained using a final wash of 1.65% Eriochrome black in distilled water.
Hemoculture
At the time of death, hemoculture in DH82 macrophages (Yabsley et al., 2004;
Hall et al., 2007) was carried out with 1 ml of EDTA-anti-coagulated whole blood and 150 checked daily for the presence of trypomastigotes. Briefly, in a 50-ml tube, approximately 35 ml of ACE lysing buffer was added to blood, gently inverted for 5 min, and centrifuged at 1,620 g for 10 min. The supernatant was discarded and the procedure repeated. The buffy coat pellet was resuspended in 5-ml of MEM and added to a confluent monolayer of DH82 cells.
Histopathology
Formalin-fixed tissues were embedded in paraffin, sectioned, and stained with hemotoxylin and eosin by standard methods. Inflammation was blindly scored based on the number of foci detected per fields viewed and compared to negative control tissues.
These scores were then evaluated and assigned to 1 of 4 categories, i.e., very mild, mild, moderate, and severe inflammation. Presence of other histologic lesions and pseudocysts were also noted.
Results
Patent infections were only detected in IV-inoculated (group 1), bug-fed (group
5), and PO-inoculated (group 6) raccoons. Parasitemias were first detected in IV- inoculated raccoons, followed by bug-fed, then PO-inoculated raccoons (Fig. 7.1). None of the negative control raccoons or raccoons that ingested amastigote-infected tissue developed parasitemias.
Trypanosoma cruzi DNA was amplified from raccoons in groups 1 (IV), 5 (bug), and 6 (PO); no T. cruzi DNA was detected by PCR in tissue-fed raccoons. The IV- inoculated raccoon was PCR-positive on day 7 post-inoculation (PI) and every bleed date thereafter. Interestingly, the first detection of T. cruzi DNA in the PO-inoculated and bug- 151 fed individuals was 1 wk later, at 14 days PI; animals remained PCR-positive through the completion of the study. The amplified product in all cases was 120bp, consistent with the lineage typing of the inoculation strain (FL RAC 9) (Roellig et al, 2008). On the day of euthanasia, for those animals that were parasitemic (groups 1, 5, and 6), T. cruzi DNA was amplified from all tissues collected. Trypanosoma cruzi DNA was not amplified from the blood or tissues of any raccoon that ingested T. cruzi-infected tissues.
Similar to the parasitemia results, seroconversion to T. cruzi only occurred in IV- inoculated, PO-inoculated, and bug-fed groups. The intravenously inoculated raccoon seroconverted sooner (7 days PI) than those that ingested infected bugs (21 days PI) or trypomastigote contaminated food (28 days PI).
After 8 wk, cultures from tissue-fed animals (groups 2-4) were considered negative for T. cruzi. Hemoculture was not performed for bug-fed animals, which were parasitemic at the time of euthanasia, but IV- and PO-inoculated groups were positive by hemoculture, confirming patent infections detected by other methods.
No major histologic lesions were noted other than varying levels of inflammation and pseudocysts in individuals that were successfully infected with T. cruzi (Table 7.1).
The IV- and PO-inoculated raccoons had greater levels of inflammation than the bug-fed raccoons, and tissue-fed raccoons had no appreciable inflammation.
Discussion
While different transmission routes for T. cruzi have been identified experimentally, the mechanism by which wildlife reservoirs predominately become infected with T. cruzi in the United States is unknown. Classic stercorarian vector 152 transmission is unlikely because the 2 main reservoirs, raccoons and opossums, rarely use permanent dens, and competent vectors are rarely found in, or near, temporary dens,
(Walton et al., 1958) and some species of native vectors, such as Triatoma sanguisuga, have delayed defecation times following the acquisition of a blood meal (Zeledón 1974).
Vertical, or transplacental, transmission has been demonstrated in rodent models
(Andrade, 1982; Moreno et al., 2003) and naturally in humans (Hoff et al., 1978, Muños et al., 2007), but similar experiments have not been performed with raccoons. Infective trypomastigote stages in breast milk have also been reported in rodent models (Miles,
1972), but experimental infection studies in rodents (Mazza et al, 1936) and opossums
(Jansen et al., 1994) disprove this as a route of transmission. Oral, or intragastric, transmission has been responsible for numerous outbreaks in humans, particularly when associated with ingesting vector-contaminated juices (Ianni and Mady, 2005).
Additionally, researchers have hypothesized this as a mode of transmission in wildlife reservoirs (Yaegar, 1971; Miles, 2004; Coura, 2006; Dias, 2006), including via the ingestion of infected bugs and other animals.
In previous IG inoculation studies with mice, researchers found that T. cruzi trypomastigotes are able to penetrate the gastric mucosa and establish infection; this process was dependent on the expression of surface proteins gp90 and gp82 (Cortez et al.,
2006; Covarrubias et al., 2007). When ingested, metacyclic trypomastigotes found in the feces of triatomes were able to produce detectable parasitemias prior to day 20 PI
(Covarrubias et al., 2007) and were more infective than blood-form trypomastigotes
(Calvo Méndez et al., 1992). Since a patent infection occurred after oral ingestion of our culture-derived trypomastigotes, we believe the parasites were more similar to 153 bloodstream-forms, and our results may simulate the indirect consequences of mesomammal carnivory by ingesting highly parasitemic, infected blood.
The oral transmission of T. cruzi via ingestion of tissues has only been presented in the literature once, but with a non-reservoir species, where Phyllostomus sp. (bats) became infected after ingesting T. cruzi- infected mice (Thomas et al., 2007). Our findings, however, suggest that raccoons, a major wildlife reservoir in the United States, do not become readily infected after consuming infected tissues. Discrepancies between our findings and those of Thomas et al. (2007) may be explained by the ingestion of bloodstream trypomastigotes. In the previous study, the bats may have become infected because the mice were highly parasitemic; in the present study, raccoons were also fed tissue from parasitemic animals. However, less infective forms may have been present in this ―inoculum‖ since the killed animals were exsanguinated at the time of death and remaining trypomastigotes may have died following the animals‘ death and clotting of remaining blood. Feeding of tissues from these animals more readily mimics natural exposure, i.e., scavenging of carcasses by wildlife, where the blood of deceased animals has clotted. Based on our results, carnivory does not appear to be a major contributor to the high prevalence of infections seen in wildlife.
As has been demonstrated with opossums and striped skunks (Yaegar, 1971;
Davis et al., 1980), raccoons develop patent infections upon ingestion of infected bugs.
The metacyclic trypomastigotes are able to withstand the acidic gastric environment and penetrate the gut mucosa to establish an infection. These findings parallel the human cases of T. cruzi resultant from ingesting vector parts in food or drink (Ianni and Mady,
2005; Shikanai-Yasuda et al., 1991), because in all cases the infective stage, metacyclics, 154 are ingested. Together, these data suggest that consumption of bugs by raccoons and opposums (both omnivorous), is the major transmission route for T. cruzi in the United
States since alternative transmission routes such as stercorarian vector transmission and ingestion of infected tissues appear to be insignificant.
Acknowledgments
The authors thank Kate McMillan, Mason Savage, Jessica Murdock, and Emily
Brown (SCWDS) for laboratory assistance. This study was supported by the National
Institutes of Health, National Institute of Allergy and Infectious Disease grant R15
AI067304.
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159
Table 7.1. Inflammatory lesions of tissues in raccoons experimentally infected with Trypanosoma cruzi via different routes. Inoculation methods were as follows: group 1 intravenously (IV) with 1 x 106 culture-derived trypomastigotes; groups 2, 3 and 4 per os with 33.3-g amastigote-infected tissue at 2, 12, and 24 hr post-mortem, respectively; group 5 per os with 3 infected R. prolixis 4th or 5th instar nymphs; and group 6 per os with 1 x 106 culture-derived trypomastigotes. Animal Tissue Skeletal Adrenal Sex LN Heart Lung Liver Spleen GI Pancreas Kidney Bladder Brain Muscle Gland Organ Group 1 (IV) RAC A Vm S † S † S Mo Vm Vm M† M † Vm M M M Group 2 (2hr) RAC B Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm RAC C Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Group 3 (12hr) RAC D Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm RAC E Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Group 4 (24hr) RAC F Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm RAC G Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Vm Group 5 (bug) RAC H Vm M Mo Vm S Vm Vm Vm Vm Vm Mo Vm Vm RAC I Vm M Mo † Vm S Vm Vm Vm Vm Vm Vm Vm Vm Group 6 (PO control) RAC J Vm Mo M Vm Mo Vm† Vm Vm S M Vm M Vm * LN= Lymph node; GI= gastrointestinal tract; Vm= very mild; M= mild; Mo= moderate, S= severe †Pseudocyst(s) found within tissue sample 160
300000 Group 1 (IV) Group 2 (2 hr) 250000 Group 3 (12 hr) Group 4 (24 hr) 200000 Group 5 (bug) Group 6 (per os)
150000
100000 Parasitemia (parasites/mL) Parasitemia 50000
0 7 DPI 14 DPI 21 DPI 28 DPI 35 DPI 42 DPI Time (days post-inoculation)
Figure 7.1. Parasitemias of raccoons experimentally infected with Trypanosoma cruzi via different inoculation methods. Inoculation methods were as follows: group 1 intravenously (IV) with 1 x 106 culture-derived trypomastigotes; groups 2, 3 and 4 per os with 33.3-g amastigote-infected tissue at 2, 12, and 24 hr post-mortem, respectively; group 5 per os with 3 infected R. prolixis 4th or 5th instar nymphs; and group 6 per os with 1 x 106 culture-derived trypomastigotes. Black arrows indicate day on which animals were euthanized.
161
CHAPTER 8
EVALUATION OF THE CHAGAS STAT-PAK ™ ASSAY FOR DETECTION OF
TRYPANOSOMA CRUZI ANTIBODIES IN WILDLIFE RESERVOIRS1
1 Roellig, D.M., Yabsley, M.J., Brown, E.L. 2008. Journal of Parasitology. 95: 775-777. Reprinted here with permission of publisher. 162
Abstract
An immunochromatographic assay (Chagas Stat-Pak™) was evaluated for the detection of Trypanosoma cruzi antibodies in 4 species of wildlife reservoirs. Antibodies to T. cruzi were detected in both raccoons (Procyon lotor) (naturally and experimentally infected) and degus (Octodon degu) (experimentally-infected) using the Chagas Stat-
Pak™. In naturally exposed wild raccoons, the Chagas Stat-Pak™ had a sensitivity and specificity of 66.7-80% and 96.3%, respectively. Compared with indirect immunofluorescent antibody assay results, seroconversion as determined by Chagas Stat-
Pak™ was delayed for experimentally infected raccoons, but occurred sooner in experimentally infected degus. The Chagas Stat-Pak™ did not detect antibodies in naturally or experimentally infected Virginia opossums (Didelphis virginiana) or in experimentally infected short-tailed opossums (Monodelphis domestica). These data suggest that the Chagas Stat-Pak™ might be useful in field studies of raccoons and degus when samples would not be available for more conventional serologic assays. Because this assay did not work on either species of marsupial, the applicability of the assay should be examined before it is used in other wild species.
Trypanosoma cruzi, the etiological agent of American trypanosomiasis or Chagas‘ disease, is an important medical and veterinary pathogen. In some hosts, such as humans and dogs, T. cruzi may cause fatal myocarditis during the chronic phase of disease.
Although an estimated 10-12 million people are infected in Latin America, autochthonous human infections in the United States are rare, with a total of only 6 cases having been reported from California, Louisiana, Texas, and Tennessee since 1955
(Woody and Woody, 1955; Navin et al., 1985; Herwaldt et al., 2000; Dorn et al., 2007). 163
Although apparently underdiagnosed, T. cruzi infection in domestic dogs has been reported from Texas, Louisiana, Oklahoma, Georgia, South Carolina, and Virginia (see
Meurs et al., 1998; Kjos et al., 2008).
In contrast, reports of T. cruzi in wildlife, e.g., raccoons and opossums, are relatively common (John and Hoppe, 1986; Yabsley et al., 2001; Brown, 2008), although disease in these reservoirs is rare. Several techniques have been utilized to determine the prevalence of T. cruzi in wildlife species, i.e., direct examination of blood, culture of blood and/or tissues, polymerase chain reaction (PCR) testing of blood and/or tissues, and serology (McKeever et al., 1958; John and Hoppe, 1986; Yabsley et al., 2001; James et al., 2002). The majority of previous studies have used hemoculture or the direct examination of blood to determine prevalence. These methods underestimate prevalence because parasitemia decreases during the chronic stage of the infection. Serological testing is considered the most sensitive assay for determining prevalence because antibodies to T. cruzi persist during the chronic phase (Yabsley et al. 2001; Yabsley and
Noblet 2002a). For example, prevalence of T. cruzi in raccoons (greater than 10 raccoons tested) from the southern United States based on culture ranged from 15-43% (Karsten et al., 1992; Pung et al., 1995; Pietrzak and Pung, 1998; Yabsley and Noblet, 2002a; Brown,
2008), while the prevalence based on serologic testing (indirect immunofluorescent antibody test, IFA) was 33-70% (Yabsley et al., 2001; Hancock et al., 2005; Brown,
2008).
Rapid immunochromatographic assays for T. cruzi have been developed and validated for use in humans and dogs (Luquetti et al. 2003; Ponce et al. 2005; Cardinal et al. 2006), but have not been validated for use in other mammalian hosts. In general, these 164 assays utilize a dye bound to an antibody-binding protein (e.g., protein A or G) which together will bind to all antibodies present in a serum, plasma, or blood sample. If antibodies specific to the recombinant antigens are present in a sample, then this protein- dye-antibody complex will be visualized as a colored band or spot. The Chagas Stat-
Pak™ (Chembio Diagnostics Inc., Medford, New York) utilizes a combination of several recombinant antigens specific for T. cruzi (antigens described by Umezawa et al., 2003).
For humans and dogs, these assays have high sensitivity and specificity and, if validated for wildlife reservoirs, would provide a field-friendly assay that is rapid, simple, stable at room temperature, and can be run with small quantities of blood, plasma, or serum. The objective of the current study was to evaluate the use of the commercially available
Chagas Stat-Pak™ for use in 4 wildlife reservoirs species, 2 from North America
(raccoon [Procyon lotor] and Virginia opossum [Didelphis virginiana]) and 2 from South
America (degu [Octodon degu] and short-tailed opossum [Monodelphis domestica]).
Archived serum samples used in this study were collected during previous studies and stored at -20 C until testing. Samples from wild hosts were collected from raccoons
(n=57) and Virginia opossums (n=11) from Georgia and Florida (Brown, 2008). Four wildlife reservoir species (raccoons, degus, Virginia opossums, and short-tailed opossums) were experimentally infected with T. cruzi to provide confirmed positive samples and to determine time to seroconversion by indirect immunofluorescent antibody
(IFA) assay and Chagas Stat-Pak™ assays. For experimental infections, raccoons, degus, and short-tailed opossums were captive bred and obtained from commercial sources and the Virginia opossums were from two litters of joeys raised in our animal facility by wild-caught nursing females (both females were IFA and culture negative for T. cruzi). 165
All animals were housed indoors and shown to be IFA and culture negative for T. cruzi before experimental inoculation. Animals were inoculated with 1 x 106 DH82 macrophage-derived trypomastigotes (Roellig et al., 2008) by intraperitoneal (degus, short-tailed opossums, Virginia opossums) or intravenous (raccoons) routes. Negative control animals were similarly inoculated with equivalent volumes of media. Blood samples were aseptically collected into ethylenediaminetetraacetic acid (EDTA) tubes at various days post-inoculation; plasma was collected and frozen at -20 C until testing
(Roellig et al., 2008; Roellig et al., unpublished). Numbers of animals sampled at each sampling date varied depending on the volume of plasma available for testing.
To confirm infections, culture attempts in LIT medium or DH82 canine macrophages were made on a subset of wild animals and on all experimental hosts as described (Yabsley and Noblet, 2002b; Yabsley et al., 2004). IFA testing of all samples was performed as described (Yabsley et al., 2001; Brown, 2008) using the following secondary antibodies: a goat-anti raccoon IgG (Kirkegaard and Perry Laboratories (KPL),
Gaithersburg, Maryland, USA) and a rabbit anti-rat IgG (KPL) for raccoons and degus, respectively. For both species of opossums, a rabbit anti-opossum IgG (Bethyl
Laboratories, Montgomery, Texas, USA) was used followed by a fluorescin-labeled anti- rabbit IgG (KPL). Slides were examined under an Olympus microscope. The Chagas
Stat-Pak™ assay was conducted per the manufacturer‘s instructions. Chi-square analysis
(p=0.05) was used to determine if differences existed between the serologic assay results.
The Chagas-Stat PakTM detected antibodies to T. cruzi in both naturally and experimentally infected raccoons, as well as experimentally infected degus (Tables 8.1,
8.2). No significant differences were noted in the prevalence of T. cruzi antibodies in 57 166 wild raccoons between IFA and Chagas Stat-Pak™ (42.1% vs. 33.3%, respectively, χ2=
0.934, p=0.3339) assays. Discordant results were obtained for 9 of 57 (15.8%) samples
(Table 8.1). We calculated the sensitivity of the Chagas Stat-Pak™ in 2 ways using data from naturally infected raccoons. First, based on a comparison with IFA positive animals only, the sensitivity of the Chagas Stat-Pak™ was 66.7% (16 of 24 IFA positives) (Table
8.1) and second, based on raccoons that were both IFA and culture positive, the sensitivity of the Chagas Stat-Pak™ was 80% (12 of 15 positives) (Table 8.1). The specificity of the Chagas Stat-Pak™ was 96.3% (based on 26 of 27 raccoons that were both IFA and culture negative). For 4 experimentally infected raccoons, the sensitivity of the Chagas Stat-Pak™ was lower compared with IFA testing during the first 3 wk of infection (Table 8.2). In contrast, the Chagas Stat-Pak™ test was more sensitive for detecting early infections in some of the experimentally infected degus (Table 8.2).
Although IFA and culture provided reliably positive results, the Chagas Stat-Pak™ failed to detect anti-T. cruzi antibodies in either experimentally-infected marsupial species we tested. Similar results were found in the naturally-infected Virginia opossums (n=7). All
IFA and culture negative opossums (4 Virginia opossums and 3 short-tailed opossums) were also negative by the Chagas‘ Stat-Pak™.
A number of different species of mammals serve as wildlife reservoir hosts for T. cruzi, and the availability of a rapid test would improve the ability to study the epizootiology of this important pathogen. Previously, a commercially-available rapid assay (Trypanosoma Detect™ MRA Rapid Test; InBios International Ltd., Seattle, WA) successfully detected T. cruzi antibodies in 2 IFA-positive gray foxes (Urocyon cinereoargenteus) from South Carolina, USA (Rosypal et al., 2007). In the present study, 167 we found that the Chagas Stat-Pak™ successfully detected antibodies in two known reservoir hosts, raccoons and degus. The assay also detected T. cruzi antibodies in 2 experimentally infected laboratory rodents (Balb/c mice and Windsor rats) (data not shown), suggesting the assay might be useful for laboratory experiments using rodents.
However, the assay failed to detect antibodies in 2 species of marsupials which is likely because the staphylococcal and streptococcal proteins commonly used in rapid tests only variably bind with antibodies from different marsupial species (Kronvall et al., 1970;
Kronvall, 1973; De Chateau et al., 1993).
The sensitivity of the Chagas Stat-Pak™ assay for naturally and experimentally infected raccoons was lower compared with studies on humans (93.4%-98.5%) (Luquetti et al., 2003; Roddy et al., 2008) and dogs (94%) (Cardinal et al., 2006). Several possible explanations could explain this finding. In the current study, we noted species differences in the sensitivity of the Chagas Stat-Pak™ during early T. cruzi infections.
Experimentally infected raccoons seroconverted, as detected by IFA, between days post- infection (DPI) 4-10; however, Chagas Stat-Pak™ seroconversion was not detected until
DPI 28, while degus seroconverted with the Chagas Stat-Pak™ by DPI 7. Previous studies did not evaluate the sensitivity of the Chagas Stat-Pak™ during acute infections
(Luquetti et al., 2003; Cardinal et al., 2006). Furthermore, previous studies of dogs and humans were conducted on banks of serum samples that were seropositive by multiple serologic assays. We only used 1 serologic assay (IFA) to determine the serostatus of the animals included in this study. Although the majority of samples are seropositive by multiple assays, discordant results between assays led to a small percentage of samples being classified as either ―borderline‖ or equivocal (Yabsley et al., 2001; Cardinal et al., 168
2006). If the samples used in our study had been tested with multiple serologic assays
(ELISA, HA, etc.), some borderline IFA positive samples might have tested negative, resulting in their exclusion and a consequent increase in assay sensitivity for that species.
These data suggest that the Chagas Stat-Pak™ might be useful in field studies of some species when samples would not be available for more conventional serologic assays or if testing is impractical. Because this assay did not work on either species of marsupial, it must be emphasized that any commercial serologic assay or rapid test must be validated for use in wild animal species before wide-spread use in epidemiological studies.
The authors would like to thank K. McMillan and the animal care staff at the
University of Georgia, College of Veterinary Medicine for assistance with experimental animals. This study was supported by the National Institutes of Health, National Institute of Allergy and Infectious Diseases grant R15 AI067304. We also would like to thank
Chembio Diagnostics Inc. for donating some of the tests used in this study.
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Table 8.1. Results of hemoculture, indirect immunofluorescent antibody (IFA), and Chagas Stat-Pak™ assay testing of 57 wild raccoons from Georgia and Florida.
Hemoculture positive Hemoculture negative Chagas Stat-Pak IFA positive IFA negative IFA positive IFA negative Positive 12 2 4 1 Negative 3 4 5 26
174
Table 8.2. Indirect immunofluorescent antibody (IFA) and Chagas Stat-Pak™ testing results for raccoons (Procyon lotor) and degus (Octodon degu) experimentally-infected with Trypanosoma cruzi.
DPI* 3 DPI 10 DPI 28 DPI 42 DPI 112‡ Negative controls IFA Stat† IFA Stat IFA Stat IFA Stat IFA Stat IFA Stat Raccoons 2/4 0/4 3/4 0/4 3/3 2/3 2/2 2/2 3/3 3/3 0/2 0/2
DPI 7 DPI 14 DPI 21 DPI 56 DPI 112‡ Negative controls IFA Stat IFA Stat IFA Stat IFA Stat IFA Stat IFA Stat Degus 0/5 2/5 6/6 3/6 4/4 3/4 2/2 2/2 3/3 1/3 0/1 0/1
*DPI, days post-inoculation. †Stat, Chagas Stat-Pak™ results ‡All animals were hemoculture positive on DPI 112 which confirmed infection in each animal.
175
CHAPTER 9
INFECTIVITY OF SYLVATIC AND DOMESTIC DOG TRYPANOSOMA CRUZI
ISOLATES FROM THE UNITED STATES TO LABORATORY MICE AND RATS1
1 Roellig, D.M. and Yabsley, M.J. Submitted to American Journal of Tropical Medicine and Hygiene, 11/04/2009. 176
Abstract
Trypanosoma cruzi, the causative agent of Chagas disease, is widespread in the southern United States. In addition to detection in numerous wildlife host species, cases have been diagnosed in domestic dogs and humans. In the current investigation, laboratory mice [Crl:CD1 (ICR)] were inoculated with 18 United States T. cruzi isolates obtained from a wide host range to elucidate their infectivity, pathogenicity, and virulence. In addition, laboratory rats (ICR strain) were inoculated with four isolates.
Mice and rats were susceptible to infection with all of the strains, but no morbidity or mortality was noted, which indicates these United States T. cruzi isolates were of low virulence for laboratory mice and rats.
Trypanosoma cruzi, the causative agent of Chagas disease, infects ~10--12 million people in the Americas, with approximately 200,000 new cases annually (1, 2). In the United States (US), only six autochthonously-acquired human infections have been reported; however, >1,000 seropositive individuals have been detected during routine screening of US blood donations since 2007 (3). While few autochthonous US human cases have been reported, reports of domestic dog and captive exotic animal cases are increasing (4, 5), and the prevalence of T. cruzi in wild mammal reservoir species can be as high as in South America (6, 7). T. cruzi is currently categorized into one of six discrete typing units (TcI, TcIIa, TcIIb, TcIIc, TcIId, TcIIe). To date, all US isolates from humans, vectors, wild mammals, domestic animals, and non-human primates have been classified as either TcI or TcIIa (8--10).
Identifying the genotype of a T. cruzi strain is often important for characterizing biological differences among isolates, such as virulence, pathogenicity, tissue tropism, 177 geographical locality, and host/reservoir capacity. Previous mouse infectivity studies of
T. cruzi from Brazil demonstrated differences due to host origin, such as sylvatic (wild) versus human or vector strains, and genotype (11, 12). Both studies reported either higher parasitemias or greater infectivity of Didelphis isolates compared with human, vector, or placental mammal isolates. Patent infections were also more frequent in laboratory mice inoculated with TcII strains compared with TcI strains (12). In contrast, US isolates rarely cause morbidity and mortality in laboratory rodents, but in a single study, a T. cruzi isolate from a raccoon caused hind limb paralysis in mice (13--21).
Differences in the outcome of infections in these previous studies may be due to use of different mouse strains, T. cruzi inoculum stage, inoculation route and/or dose, and source (host) species of the isolate. Additionally, many studies were conducted with genetically unclassified strains. The goal of the current study was to experimentally infect laboratory mice with genetically classified US T. cruzi isolates from a wide host range to determine infectivity, pathogenicity, and virulence. Based on previous studies on US strains, we hypothesized that sylvatic isolates would be infective but not virulent to mice.
A total of 18 T. cruzi isolates from seven mammalian host species and two vector species was used in the study (Table 9.1). These isolates were chosen to represent both genotypes (TcI and TcIIa) and a diverse geographic and host range. Two isolates from
Brazil (Brazil and Y strains) were used as positive controls [kindly provided by Dr. Rick
Tarleton (University of Georgia, Athens, GA)]. Parasites stored in liquid nitrogen (first passage for all but the two Brazil strains) were rapidly thawed and established in DH82 canine macrophage monolayers to yield the infective culture-derived trypomastigotes
(22). 178
One hundred and eighty-two outbred 8-wk-old male Crl:CD1 (ICR) mice and 16 white laboratory rats (Charles River Laboratory International, Inc, Wilmington, MA) were housed in microisolator cages in climate-controlled animal facilities at the College of Veterinary Medicine, University of Georgia (Athens, GA). All methods were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of
Georgia. Mice were weighed and randomly separated into one of 21 groups (18 US isolate groups, two positive control groups, and one negative control group); rats were weighed and separated into one of five groups (four US isolates groups and one negative control). Individuals (mouse, n=9; rat, n=4) from each experimental and positive control group were inoculated intraperitoneally with 1 x 106 culture-derived trypomastigotes of one of the representative strains (Table 9.1). Negative controls (n=2) were similarly inoculated with an equivalent volume of culture medium. Any physical or behavioral changes indicative of Chagas clinical signs, such as lethargy, hind limb paralysis, weight loss, or ruffed coat, were noted daily. At days 3, 7, 10, 14, 17, 21, 24, 28, and 112 post- inoculation (DPI), one mouse from each experimental and positive control group was humanely euthanized via CO2 and exsanguination. At days 3, 7, 28, and 112 post- inoculation, one rat from each experimental group was humanely euthanized. At euthanasia, approximately 1 to 1.5 mL of whole blood from mice and 3 mL of whole blood from rats were collected by cardiac puncture into ethylenediaminetetraacetic acid
(EDTA) tubes. All animals were weighed prior to euthanasia at the above time points.
Infection was determined by detecting T. cruzi DNA by polymerase chain reaction (PCR) or culture of blood. For PCR, DNA was extracted from 100 µl of whole blood and sections of heart and quadriceps muscle collected at necropsy. Amplification 179 of the 24Sα rDNA gene of T. cruzi using a modified nested reaction was performed as previously described (22--24). For culture, remaining whole blood was centrifuged at
1620 x g for 15 min. Plasma was removed and approximately 3 mL of liver infusion tryptose (LIT) medium added to the remaining buffy coat and red blood cells (25).
Cultures were evaluated between 2 months and 4 months.
Based on PCR and culture results, all eighteen US isolates caused patent infections in mice (Table 9.1). Results from the first eight bleeding periods (3 to 28 DPI) were combined as a measure of infection status during the acute stage. During the acute stage, at least one sample from at least one animal was positive either by hemoculture or
PCR but only a single isolate (Brazil) caused infections in each mouse at each time period. Overall, during the acute stage, significantly more mice inoculated with US TcI strains were parasitemic (as determined by PCR assay of blood) with T. cruzi compared with mice inoculated with TcIIa strains (F=4.9532, p=0.043). Additionally, significant differences in T. cruzi present in tissue (heart and/or quadriceps) between genotype were noted. More US origin TcI-inoculated mice tested T. cruzi-positive in all tissues compared with TcIIa-inoculated mice (F=5.1317, p=0.0399); however, no difference was noted in tissue predilection (heart vs. quadriceps muscle) between the two genotypes
(F=1.5217, p=0.2377).
Infectivity for mice was not strictly associated with host origin or genotype. For example, two TcI isolates from Virginia opossums (GA Opo 75 and GA Opo 43) yielded dramatically different results with many more mice having detectable infections with the
GA Opo 75 strain compared with GA Opo 43 (Table 9.1). Mortality and weight loss were not observed in individuals inoculated with any US isolates of T. cruzi (data not shown); 180 however, one mouse inoculated with the Y-strain had marked weight loss and lethargy and was humanely euthanized. Parasites were detected in the blood and tissues of this animal.
Similarly, all four isolates caused patent infections in laboratory rats(Table 9.2).
T. cruzi was detected in the blood and tissues of rats infected with both US TcI isolates on multiple occasions and one rat was parasitemic at 112 DPI (USA Opossum). No rats inoculated with TcIIa isolates had detectable parasitemias by PCR but an animal from each TcIIa group was culture positive on 3 DPI. Although infection differences were noted between genotypes, small sample sizes and few isolates precluded statistical inferences.
In this study, the two T. cruzi genotypes caused differential infection dynamics in mice and rats as determined by PCR detection of T. cruzi DNA in the blood and tissues.
In general, significantly more TcI-inoculated mice had detectable infections compared with TcIIa-inoculated mice. Additionally, chronic mice and rats inoculated with two TcI isolates (FL Opo 18 and/or USA Opossum) maintained parasitemias as indicated by PCR assay of blood, while no TcIIa-inoculated mice or rats were parasitemic at the chronic stage of infection. These findings suggest that the US sylvatic TcI isolates in this study had greater infectivity to laboratory rodents compared with US TcIIa isolates. Previous studies of US T. cruzi isolates have not investigated infectivity differences based on genotype; however, a study of T. cruzi isolates from Brazil found TcII strains more infective and virulent than TcI strains (12). T. cruzi from Brazil and all South American countries is markedly different from US T. cruzi, particularly TcII strains as all TcII subtypes are found in South America while only TcIIa has been detected in the US (26; 181
Roellig et al., unpublished). Additionally, US TcIIa strains are genetically distinct from
South American TcIIa strains at numerous loci (27; Roellig et al., unpublished). It is possible that these molecular differences between T. cruzi from South America and the
United States may account for biological differences, including infectivity to mice and rats.
We also noted in this study that none of the laboratory rodents inoculated with US strains resulted in morbidity or mortality. These data support previous studies which reported that US sylvatic isolates rarely produced detectable infections or produced intermittent parasitemias without morbidity or mortality (13—19, 21). In contrast, T. cruzi isolates from South America readily infect a wide variety of laboratory mice strains and many cause significant morbidity and mortality (11, 12). In the current study, one control mouse inoculated with Y strain displayed lethargy and marked weight loss.
Although no clinical signs were observed in mice inoculated with the Brazil strain in this study, this strain has previously been shown to cause disease and mortality (28). A single previous study observed mortality in three of four C3H mice inoculated with a US T. cruzi isolate from a North Carolina raccoon; these mice were not parasitemic but amastigotes were observed in muscle tissue (20). Further, natural infections of captive exotic animals have resulted in mortality indicating that some strains of T. cruzi from the
US can cause disease and death (5). These data combined suggest that the biological characteristics of sylvatic US T. cruzi isolates may vary considerably. 182
Acknowledgements
The authors thank B. Shock, E.L. Blizzard, and E. Gleim for laboratory assistance and the Animal Resource staff at The University of Georgia College of Veterinary Medicine for assistance with mouse care.
Financial Support
This study was primarily supported by the National Institutes of Health, National
Institute of Allergy and Infectious Disease Grant R15 AI067304. Additional support was through funding provided to SCWDS by the Federal Aid to Wildlife Restoration Act (50
Stat. 917) and through sponsorship of the fish and wildlife agencies of Alabama,
Arkansas, Florida, Georgia, Kansas, Kentucky, Louisiana, Maryland, Mississippi,
Missouri, North Caroling, Oklahoma, Puerto Rico, South Carolina, Tennessee, Virginia, and West Virginia.
Authors‘ Addresses: Dawn M. Roellig and Michael J. Yabsley, 589 D.W. Brooks Drive,
Wildlife Health Building, College of Veterinary Medicine, The University of Georgia,
Athens, Georgia 30602, Telephone: 706-542-1741, Fax: 706-542-5865, E-mails: [email protected] and [email protected].
183
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Table 9.1. Detection of Trypanosoma cruzi in eight acutely-infected and one chronically-infected Crl:CD1 (ICR) mice.
No. of mice that were PCR No. positive in acute stage (+, chronic Hemoculture mouse was positive) positive Isolate Host Origin Lineage* Blood Heart Quadriceps muscle FL Opo 18 Didelphis virginiana Wakulla Co., FL I 4 (+) 1 7 1 FL Opo 3 D. virginiana Wakulla Co., FL I 5 2 6 (+) 1 USA Opossum D. virginiana Orleans Parish, LA I 6 (+) 5 (+) 7 (+) 0 GA Opo 75 D. virginiana Clarke Co., GA I 6 7 (+) 7 8 GA Opo 43 D. virginiana Chatham Co., GA I 2 0 0 1 TxTg2 Triatoma gerstackeri TX I 5 3 (+) 7 (+) 0 Florida C16 T. sanguisuga Alacua Co., FL I 5 0 6 (+) 3 TX WR 22 Neotoma micropus Uvalde Co., TX I 5 7 6 4 TX WR 30 N. micropus Uvalde Co., TX IIa 5 1 (+) 6 (+) 1 FL Rac 9 Procyon lotor Liberty Co., FL IIa 2 4 3 1 TX08 Rac 5 P. lotor Uvalde Co., TX IIa 5 2 (+) 7 (+) 2 FL Rac 13 P. lotor Leon Co., FL I/IIa 5 2 (+) 7 (+) 2 OK Dog Canis familiaris Osage and Washington Cos., IIa 3 5 7 1 OK Griffin Dog C. familiaris Coffee Co., TN I/IIa 6 1 8 (+) 4 Clarence RTL Lemur catta Liberty Co., GA IIa 2 2 3 0 RTL Meg L. catta Liberty Co., GA IIa 3 0 3 (+) 2 GA Sk 1 Mephitis mephitis Long Co., GA IIa 4 0 6 2 GA Arm 20 Dasypus novemcinctus Chatham Co., GA IIa 3 1 4 0 Brazil Human Brazil I 8 8 8 (+) 5 Y Human Brazil IIb 6 8 8 2 *As previously determined in 22 or following previously described methodology in 23. 188
Table 9.2. Detection of Trypanosoma cruzi in three acutely-infected and one chronically-infected white laboratory mice.
No. of rats that were PCR positive No. in acute stage (+, chronic rat was Hemoculture positive) positive Isolate Host Origin Lineage* Blood Heart Quadriceps muscle USA Opossum D. virginiana Orleans Parish, LA I 3 (+) 3 2 2 Florida C16 T. sanguisuga Alacua Co., FL I 3 2 (+) 3 (+) 2 TX WR 30 N. micropus Uvalde Co., TX IIa 0 0 0 1 GA Sk 1 Mephitis mephitis Long Co., GA IIa 0 0 2 1 *As previously determined in 22 or following previously described methodology in 23. 189
CHAPTER 10
CONCLUSIONS
The ultimate goal of this dissertation was to identify molecular and biological characteristics of Trypanosoma cruzi in the United States (US) and associate such characters with infection dynamics and virulence in mammalian hosts. T. cruzi is a flagellated protozoan parasite endemic to the Americas and the causative agent of
Chagas‘ disease, a major vector-borne, zoonotic disease in Central and South America.
With new human cases, increasing numbers of veterinary cases, and influx of potentially infected immigrants, understanding the ecology in the United States is imperative.
Study 1 (Chapter 3)
In contrast to studies conducted on South American isolates where six genotypes of T. cruzi have been identified, only two genotypes (TcI and TcIIa) were identified in the current study. These data support investigations in Central America and Mexico that have found a paucity of genotypes (Bosseno et al., 2002; Iwagami et al., 2007).
Additionally, further evidence for correlations between host specificity and genotype of
T.cruzi was revealed. T. cruzi isolated from humans and marsupials in the US are typically TcI, while raccoons, skunks, domestic dogs, and prosimians are typically infected with TcIIa. Although we only detected TcI in triatomid bugs, previous studies have detected TcIIa in triatomids from the US (Barnabé et al., 2001). 190
Study 2 (Chapter 4)
In the current investigation, genetic diversity was demonstrated among T. cruzi isolates from the United States. Multi-locus sequencing (Tc52, MSH2, DHFR-TS, and
COII-ND1) confirmed that US T. cruzi samples had sequence identity to TcI or TcIIa strains frm other geographic regions. Regardless of gene target, TcIIa isolates were clearly distinguished from the South American TcIIa reference strain which provided additional evidence for considerable divergence within this lineage (Machado and Ayala,
2001; Westenberger et al., 2005). Heterogeneity was also demonstrated with the identification of single nucleotide polymorphisms in sequences of Tc52 and MSH2 genes, some of which were non-synonymous changes. These changes in amino acids may account for virulence differences between strains due to changes in GSH binding efficacy, in the case of Tc52, or differences in genome maintenance, in the case of MSH2
Phylogenies of DHFR-TS (nuclear) and COII-ND1 (mitochondrial) revealed genealogical relationships among isolates and illustrate evidence for genetic exchange.
The nuclear phylogeny of DHFR-TS exhibited four major clades, three consisting of TcII strains and one TcI. Isolates of TcI from the US clustered with S. American isolates, illustrating the limited genetic variability of the lineage reported in previous studies with this gene (Machado and Ayala, 2001; Iwagami et al., 2007). More variation was noted for
TcII isolates with the US TcII clade being separate from the S. American TcIIa isolates.
The phylogeny of COII-ND1 demonstrated greater genetic diversity with additional clustering occurring within the four clades present. As with the nuclear phylogeny, TcIIa strains from the US diverged from TcII strains of S. America. The most compelling finding from the COII-ND1 phylogeny is the clustering of several TcI strains (classified 191 based on several nuclear genes) within the US TcIIa clade. The incongruence of nuclear and mitochondrial phylogenies for these isolates illustrated evidence of rare genetic exchange events in the US T. cruzi population.
Study 3 (Chapter 5)
In the current study we determined T. cruzi infection dynamics in two common
North American reservoirs, Virginia opossums (Didelphis virginiana) and raccoons
(Procyon lotor) inoculated with TcI, TcIIa, TcIIb, and dual (TcI and TcIIa) T. cruzi.
Differences in infectivity of certain genotypes to specific hosts were observed. In
Virginia opossums, only those inoculated with TcI maintained parasitemias, while those inoculated with TcIIa or TcIIb had intermittent to undetectable infections. While raccoons were able to develop a patent infection when inoculated with TcI and TcII, the highest parasitemia from a single isolate inoculation was seen with TcIIa., suggesting raccoons are better hosts for TcIIa. These data suggest that infection dynamics of various
T. cruzi strains can differ considerably in different wildlife hosts, and infectivity of different genotypes to N. American marsupials and placental mammals varies. The current study also provides evidence that a native wildlife reservoir of the United States can develop an infection with a non-native strain of T. cruzi. The ability of raccoons to serve as a potential reservoir for non-native strains of T. cruzi has obvious medical and veterinary implications.
192
Study 4 (Chapter 6)
This study evaluated infection dynamics of four genetically and geographically diverse T. cruzi strains in two South American reservoirs, degus (Octodon degus) and gray short-tailed opossums (Monodelphis domestica). Our findings suggest that short- tailed opossums may serve as reservoirs for multiple strains of T. cruzi, including TcI,
TcIIb, TcIIe, and mixed infections. These data expand our knowledge on the genotypes to which gray short-tailed opossums are susceptible, which were previously limited to TcIIc and TcI based on natural infections (Yeo et al., 2005; Roque et al., 2008). Degus developed patent infections after inoculation with each of the four isolates (representing genotypes TcI, TcIIa, TcIIb, and TcIIe) analyzed in this study. These data support molecular typing studies conducted on isolates from naturally-infected degus from South
America that showed natural infection with multiple genotypes including TcI, TcIIa,
TcIIb, TcIId, and TcIIe singly, and some mixed infections (Yeo et al., 2005; Campos et al., 2007; Rozas et al., 2007; Spotorno et al., 2008). The current study also demonstrated that degus and short-tailed opossums are competent hosts for strains of T. cruzi from
North America and no differences in infectivity based on the geographical origin of the isolates were observed.
Study 5 (Chapter 7)
This study is the first demonstration of oral transmission of Trypanosoma cruzi to raccoons, a natural reservoir host in the United States, by ingestion of trypomastigotes and infected bugs, but not infected tissue since patent infections could not be detected.
Numerous claims have been made about the importance of carnivory in maintenance of 193 the sylvatic cycle (see Dias, 2006; Miles, 2004; Coura 2006), our findings, however, suggest that raccoons do not become readily infected after consuming infected tissues. As has been demonstrated with opossums and striped skunks (Yaegar, 1971; Davis et al,
1980), raccoons develop patent infections upon ingestion of infected bugs. These findings parallel the human cases of T. cruzi resultant from ingesting vector parts in food or drink
(Ianni and Mady, 2005; Shikanai-Yasuda et al, 1991), because in all cases the infective stage, metacyclics, are ingested. Together, these data suggest that consumption of bugs by raccoons and opossums (both omnivorous), is the major transmission route for T. cruzi in the United States since alternative transmission routes such as stercorarian vector transmission and ingestion of infected meats appear to be insignificant.
Study 6 (Chapter 8)
In the present study, we found that the Chagas Stat-Pak™ successfully detected antibodies in two known reservoir hosts, raccoons and degus. The assay also detected T. cruzi antibodies in 2 experimentally infected laboratory rodents (Balb/c mice and
Windsor rats), suggesting the assay might be useful for laboratory experiments using rodents. However, the assay failed to detect antibodies in 2 species of marsupials which is likely because the staphylococcal and streptococcal proteins commonly used in rapid tests only variably bind with antibodies from different marsupial species (Kronvall et al.,
1970; Kronvall, 1973; De Chateau et al., 1993). These data suggest that the Chagas Stat-
Pak™ might be useful in field studies of some species when samples would not be available for more conventional serologic assays or if testing is impractical. Because this assay did not work on either species of marsupial, it must be emphasized that any 194 commercial serologic assay or rapid test must be validated for use in wild animal species before wide-spread use in epidemiological studies.
Study 7 (Chapter 9)
In this study, the two genotypes of US T. cruzi caused differential infection dynamics in mice and rats as determined by PCR detection of T. cruzi DNA in the blood and tissues. In general, significantly more TcI-inoculated mice had detectable infections compared with TcIIa-inoculated mice. Additionally, chronic mice and rats inoculated with two TcI isolates (FL Opo 18 and/or USA Opossum) maintained parasitemias as indicated by PCR assay of blood, while no TcIIa-inoculated mice or rats were parasitemic at the chronic stage of infection. These findings suggest that the US sylvatic TcI isolates in this study had greater infectivity to laboratory rodents compared with US TcIIa isolates. In contrast to South American T. cruzi strains, US isolates did not cause morbidity or mortality in laboratory rodents, suggesting decreased virulence of sylvatic isolates from the US. These data combined suggest that the biological characteristics of sylvatic US T. cruzi isolates may vary considerably.
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