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Toxoplasma gondii in
Australian marsupials
Nevi Parameswaran
Bachelor of Science in Veterinary Biology. Murdoch University
Bachelor of Veterinary Medicine and Surgery. Murdoch University
Faculty of Health Sciences
School of Veterinary and Biomedical Sciences
Murdoch University
Western Australia
This thesis is presented for the degree of Doctor of Philosophy of Murdoch University, 2008
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I declare that this thesis is my own account of my research and contains as its main content
work which has not previously been submitted for a degree at any tertiary education
institution.
....................................
Nevi Parameswaran
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Abstract
Diagnostic tools were developed and utilised to detect Toxoplasma gondii infection in a
range of Australian marsupial species and identify epidemiological trends. An ELISA was
developed to detect anti-T. gondii IgG in macropod marsupials. When compared with the
commercially available MAT (modified agglutination test), the ELISA was in high
agreement and yielded a κ coefficient of 0.96. Of 18 western grey kangaroos (Macropus
fuliginosus) tested for the presence of T. gondii DNA by PCR, the 9 ELISA positive
kangaroos tested PCR positive and the 9 ELISA negative kangaroos tested PCR negative
indicating that the ELISA protocol was both highly specific and sensitive and correlated
100% with the more labour intensive PCR assay.
A T. gondii seroprevalence study was undertaken on free ranging Australian marsupials.
There was a T. gondii seroprevalence of 15.5% (95%CI: 10.7-20.3) in western grey
kangaroos located in the Perth metropolitan area. The T. gondii seroprevalence in male
western grey kangaroos was significantly less than their female counterparts (p=0.038),
which may be related to behavioural differences causing differences in exposure to oocysts
or recrudescence of T. gondii infection in pregnant females. Marsupial populations located
in islands free from felids had a low overall T. gondii seroprevalence. A case control study
determined that marsupials located in areas where felids may roam are 14.20 (95%CI: 1.94-
103.66) times more likely to be T. gondii seropositive than marsupials located on felid-free
islands.
PCR, immunohistochemistry and serological techniques were used to detect T. gondii
infection in marsupial dams and their offspring. T. gondii DNA was detected in the pouch
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young of chronically infected western grey kangaroos and a woylie (Bettongia penicillata).
T. gondii DNA was also identified in the mammary gland of the woylie dam suggesting that
infection of the woylie pouch young was from suckling milk from the mammary gland.
Results of the study demonstrate that vertical transmission of T. gondii occurs in Australian
marsupials and may be of importance in the maintenance of T. gondii infection in Australian
marsupial populations.
Animal tissue and meat from Australia, predominately from Australian marsupials, were
screened for T. gondii DNA using PCR primers for the multi-copy, T. gondii specific B1
gene. Sequencing of the B1 gene revealed atypical genotypes in 7 out of 13 samples from
Australia. These 7 isolates contained single nucleotide polymorphisms (SNPs) in the B1
gene that could not be matched with known sequences from strains I, II, III and X. Six
unique genotypes were identified out of the 7 atypical isolates; two out of the 7 isolates had
the same unique sequence at the B1 gene whereas the other 5 isolates each had different
combinations of SNPs at the B1 gene. A majority of T. gondii isolates sampled from native
Australian marsupials were of an atypical genotype. The discovery of atypical strains of T.
gondii in Australia leads to further questions regarding the origin and transmission of these
atypical strains. Additional studies linking atypical strains with their clinical manifestation
are also warranted.
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Table of ContentsAbstract ............................................................................................................................ iiiTable of Contents .............................................................................................................. vList of Tables................................................................................................................... viiList of Figures ................................................................................................................ viiiList of Abbreviations........................................................................................................ ixPublications ....................................................................................................................... xConferences Abstracts....................................................................................................... xAcknowledgements ......................................................................................................... xii1. General introduction.................................................................................................. 1
1.1. Toxoplasma gondii ............................................................................................ 11.2. Life cycle........................................................................................................... 2
1.2.1. Introduction ............................................................................................... 21.2.2. Oocyst transmission .................................................................................. 31.2.3. Bradyzoite transmission ............................................................................ 51.2.4. Vertical transmission................................................................................. 5
1.3. Pathogenesis and immunity .............................................................................. 81.4. Diagnosis......................................................................................................... 101.5. Molecular epidemiology of T. gondii ............................................................. 14
1.5.1. Introduction ............................................................................................. 141.5.2. Clonal lineages of T. gondii .................................................................... 151.5.3. The effect of T. gondii genotype on disease manifestation..................... 171.5.4. Atypical T. gondii genotypes .................................................................. 20
1.6. Significance of T. gondii in Australian marsupials......................................... 231.6.1. Life cycle of T. gondii in Australian marsupials..................................... 241.6.2. T. gondii associated disease in Australian marsupials ............................ 271.6.3. Diagnosis of T. gondii infection in Australian marsupials...................... 301.6.4. Prevalence of T. gondii in Australian marsupials ................................... 33
1.7. Aims of this thesis........................................................................................... 341.8. Study design .................................................................................................... 36
2. The development of an in-house ELISA for the detection of anti-T. gondiiantibodies in macropod marsupials................................................................................. 38
2.1. Introduction ..................................................................................................... 382.2. Materials and methods .................................................................................... 39
2.2.1. Sample collection .................................................................................... 392.2.2. Modified agglutination test ..................................................................... 402.2.3. Cell culture of T. gondii tachyzoites ....................................................... 412.2.4. ELISA development................................................................................ 422.2.5. ELISA validation .................................................................................... 442.2.6. DNA extraction ....................................................................................... 452.2.7. PCR ......................................................................................................... 462.2.8. Statistics .................................................................................................. 49
2.3. Results ............................................................................................................. 492.4. Discussion ....................................................................................................... 50
3. Seroprevalence of T. gondii in free ranging Australian marsupials........................ 573.1. Introduction ..................................................................................................... 57
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3.2. Materials and methods .................................................................................... 613.2.1. Western grey and eastern grey kangaroos............................................... 613.2.2. Woylies ................................................................................................... 613.2.3. Marsupials and native rodents in island populations .............................. 623.2.4. Chuditch .................................................................................................. 633.2.5. Statistics .................................................................................................. 63
3.3. Results ............................................................................................................. 643.4. Discussion ....................................................................................................... 65
4. Vertical transmission of T. gondii in Australian marsupials.................................. 774.1. Introduction ..................................................................................................... 774.2. Materials and methods .................................................................................... 80
4.2.1. Sample collection .................................................................................... 804.2.2. Serology .................................................................................................. 814.2.3. Immunoblotting....................................................................................... 824.2.4. DNA extraction and PCR........................................................................ 834.2.5. Histology and immunohistochemistry .................................................... 84
4.3. Results ............................................................................................................. 844.3.1. Serology .................................................................................................. 844.3.2. Immunoblotting....................................................................................... 854.3.3. PCR ......................................................................................................... 864.3.4. Histology and immunohistochemistry .................................................... 87
4.4. Discussion ....................................................................................................... 875. Molecular characterization of T. gondii isolates from Australia.......................... 100
5.1. Introduction ................................................................................................... 1005.2. Materials and methods .................................................................................. 102
5.2.1. Sample collection .................................................................................. 1025.2.2. DNA extraction and PCR...................................................................... 1045.2.3. DNA sequencing ................................................................................... 104
5.3. Results ........................................................................................................... 1055.3.1. PCR of the B1 gene and sequencing of PCR products ......................... 1055.3.2. Clinical history and pathology of PCR positive animals ...................... 106
5.4. Discussion ..................................................................................................... 1086. General discussion ................................................................................................ 122
6.1. Introduction ................................................................................................... 1226.2. Diagnosis of T. gondii infection in Australian marsupials............................ 1226.3. Epidemiology of T. gondii in Australian marsupials .................................... 1276.4. Suggestions for future research..................................................................... 1356.5. Concluding remarks ...................................................................................... 137
References ..................................................................................................................... 139
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List of Tables
Table 2.1Level of agreement between a commercially available MAT and an ELISA in western greykangaroos, eastern grey kangaroos and agile wallabies ........................................................52Table 2.2PCR results of ELISA positive and negative western grey kangaroos (Group B) ................53Table 3.1Prevalence of anti-T. gondii IgG in western grey kangaroos in Perth, WA as determined byan ELISA ...............................................................................................................................71Table 3.2Prevalence of anti-T. gondii IgG in eastern grey kangaroos as determined by an ELISA ....71Table 3.3Prevalence of anti-T. gondii IgG in woylies in Australia as determined by the MAT ..........72Table 3.4Prevalence of anti-T. gondii IgG in animals in Faure Island as determined by the MAT.....72Table 3.5Prevalence of anti-T. gondii IgG in animals in Barrow Island as determined by the MAT..73Table 3.6Combined data of anti-T. gondii IgG in marsupials located in areas where cats may roam .73Table 3.7Combined data of anti-T. gondii IgG in marsupials located in areas without cats ................74Table 3.8The effect of being located in an area where cats may roam on T. gondii seropositivity inAustralian marsupials ............................................................................................................75Table 4.1MAT results from agile wallabies and their offspring...........................................................94Table 4.2ELISA and PCR results from western grey kangaroo dams and their pouch young.............95Table 4.3T. gondii DAT and MAT titres of seropositive western grey kangaroo dams ......................96Table 4.4T. gondii DAT and MAT titres of seropositive agile wallaby dams .....................................96Table 4.5T. gondii PCR results of a woylie dam and its pouch young.................................................96Table 5.1Tissue samples tested for T. gondii DNA using PCR of the B1 gene .................................113Table 5.2Summary of polymorphisms in the B1 gene from Australian T. gondii isolates ................121
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List of Figures
Figure 1.1. Major routes of transmission of T. gondii.......................................................3Figure 2.1. Checker board system to determine initial serum and antigen dilutions for anELISA ............................................................................................................................. 54Figure 2.2. Checker board system to determine secondary and tertiary dilutions for anELISA ............................................................................................................................. 55Figure 2.3. Checker board system to determine final serum dilution for an ELISA ...... 56Figure 3.1. Locations of marsupials sampled for anti-T. gondii IgG in Australia ..........76Figure 4.1 Comparative immunoblots of seropositive agile wallaby dams and theiryoung............................................................................................................................... 97Figure 4.2 Comparative immunoblots of seropositive western grey kangaroo dams andtheir pouch young............................................................................................................ 98Figure 4.3 Non-nested B1 PCR of western grey kangaroo tissue DNA ......................... 99Figure 4.4 Nested B1 PCR of western grey kangaroo tissue DNA ................................ 99
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List of Abbreviations
CI Confidence interval
DAT Direct agglutination test
DEC Department of Environment and Conservation
DNA Deoxyribonucleic acid
ELISA Enzyme linked immunosorbent assay
HCl Hydrochloric acid
IFAT Indirect fluorescent antibody test
MAT Modified agglutination test
NaCl Sodium chloride
OD Optical density
OR Odds ratio
PBS Phosphate buffered saline
PCR Polymerase chain reaction
PCR-RFLP PCR with restriction fragment length polymorphism
PY Pouch young
SA South Australia
SD Standard deviation
NSW New South Wales
WA Western Australia
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Publications
Parameswaran, N., O’Handley, RM., Vitali, S., Walton, S., Fenwick, SG., Thompson, RCA.(2006) The investigation of vertical transmission of Toxoplasma gondii in zoo bredmarsupials, 2006 ARAZPA (Australasian Regional Association of Zoological Parks andAquaria) Conference Proceedings. Australia. www.arazpa.org.au
Thompson, RCA., Traub, RJ. and Parameswaran, N. (2007) Molecular Epidemiology ofFood-borne Parasitic Zoonoses. In: Food-Borne Parasitic Zoonoses (K.D. Murrell and B.Fried Eds). Springer
Parameswaran, N, Wayne, A. Thompson, RCA. (2008) Toxoplasma. Progress report of theWoylie Conservation Research Project: diagnosis of recent woylie (Bettongia penicillataogilbyi) declines in southwestern Australia: a report to the Department of Environment andConservation Corporate Executive. http://www.naturebase.net/content/view/3230/1/1/6/ (edA Wayne). Department of Environment and Conservation, Kensington, WA. pp. 237–245
Parameswaran, N., O’Handley, RM., Grigg, ME., Fenwick, SG., Thompson, RCA. (2009)Seroprevalence of Toxoplasma gondii in wild kangaroos using an ELISA. ParasitologyInternational, 58, 161-165
Parameswaran, N., O’Handley, RM., Grigg, ME., Wayne, A., Thompson, RCA. (2009)Vertical transmission of Toxoplasma gondii in Australian Marsupials. Parasitology(manuscript accepted for publication April 2009)
Conferences Abstracts
Parameswaran, N., O’Handley, RM., Vitali, S., Fenwick, SG., Thompson, RCA. (2005)Toxoplasma gondii in Marsupials: The Utilisation of Zoo Records to Track Patterns ofVertical Transmission, WAAVP (World Association for the Advancement of VeterinaryParasitology) Conference, October 2005, Christchurch, New Zealand- Poster presentation
Parameswaran, N., O’Handley, RM., Vitali, S., Walton, S., Fenwick, SG., Thompson, RCA.(2006) The Investigation of Vertical Transmission of Toxoplasma gondii in Zoo BredMarsupials. ARAZPA (The Australasian Regional Association of Zoological Parks andAquaria) Conference. March 2006. Perth. Australia. Oral presentation
Parameswaran, N., O’Handley, RM., Vitali, S., Walton, S., Fenwick, SG., Thompson, RCA.(2006) The Investigation of Vertical Transmission of Toxoplasma gondii in Marsupials.ASP (Australian Society of Parasitology) Conference. July. Gold Coast. Australia. Oralpresentation
Parameswaran, N., O’Handley, RM., Vitali, S., Walton, S., Fenwick, SG., Thompson, RCA.(2006) The Investigation of Vertical Transmission of Toxoplasma gondii in Marsupials.ICOPA (International Congress of Parasitology) Conference. August. Glasgow. Scotland.Poster presentation
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Parameswaran, N., O’Handley, RM., Thompson, RCA. (2007) Development of an ELISAfor the detection of Toxoplasma gondii antibodies in macropod marsupials. ASP (AustralianSociety of Parasitology) Conference. July. Canberra. Australia. Oral presentation
Parameswaran, N., O’Handley, RM., Thompson, RCA. (2007) Development of an ELISAfor the detection of Toxoplasma gondii antibodies in macropod marsupials. WAAVP(World Association for the Advancement of Veterinary Parasitology) Conference. August.Gent. Belgium. Poster presentation
Parameswaran, N., O’Handley, RM., Thompson, RCA. (2007) Toxoplasma in AustralianMarsupials: Analysis of pouch young of naturally infected wild kangaroos for evidence ofvertical transmission. WAAVP (World Association for the Advancement of VeterinaryParasitology) Conference. August. Gent Belgium. Oral presentation
Parameswaran, N., Grigg, ME., O’Handley, RM., Thompson, RCA. (2008) Geneticdiversity among Australian isolates of Toxoplasma gondii. ASP (Australian Society ofParasitology) Conference. July. Glenelg. Australia. Oral and poster presentation
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Acknowledgements
I would not have been able to get through this PhD project without the help and support of
my family, friends and colleagues. Firstly, I would like to thank my supervisors, Prof
Andrew Thompson, Dr Ryan O’Handley and A/Prof Stan Fenwick for believing in me and
for their guidance and encouragement through both the ups and downs of this project.
Many people helped me find my feet in the laboratory environment and taught me
techniques and principles I will continue to use throughout my research career. Ian
Roberson kindly answered all my epidemiological questions throughout the project. Peter
Adams was considerate and patient enough to introduce me to the ins and outs of PCR.
Ryan O’Handley, Rob Steuart and Andrew Mikosa not only spent several selfless hours
teaching me many different forms of lab work, they also gave me friendship, laughter and a
shoulder. I must also acknowledge other lab buddies who were there for me from the
beginning and have since moved location, but who continue to be a source of warmth and
intuitive advice- Rebecca Traub, Carly Palmer and Zablon Njiru.
I was fortunate enough to do fieldwork in a number of beautiful locations, with a number of
kind and fun loving people. Glen Goudie and his kangaroo shooting team were gentle giants
and helped me get the samples I needed to make this project work. Chris Mayberry and Lisa
Hulme-Moir were of immense assistance in kangaroo sampling and sample processing and I
am deeply indebted to them. Adrian Wayne and his team at the Department of Environment
and Conservation, Manjimup, gave me wonderful fieldwork experience and all the woylie
blood I could ever need. I also had a fantastic and productive time doing fieldwork with
Andy Smith, who also kindly provided me with blood samples from a range of weird and
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wonderful Australian wildlife. I would also like to thank Simon Walton for his help and
enthusiasm in the provision of agile wallaby blood and data. I also collaborated with Tamsin
Barnes and Michael Roberts, who provided me with sera from kangaroos on the east coast
of Australia that I would never have been able to obtain on my own. Simon Vitali, Paul
Eden and several other staff at Perth Zoo were extremely supportive throughout my PhD
project and gave me data, contacts, samples and recent news on anything related to
Toxoplasma at the zoo.
I don’t know what I would have done without the amazing support of my mother and sister
throughout this project. I would like to thank my mother, who selflessly provided me with
emotional and financial support, hot meals and affection. I would like to thank my sister for
always being there for me when I needed a friend to turn to for encouragement and a
listening ear. My thanks also go to Kyne, for putting up with me in the final stages of my
PhD, giving me warmth and helping stay me positive. I would also like to thank my very
close friends Anusha, Ghirijha, Sharon, Tabita and Girisha for giving me perspective, a
social life and plenty of distractions.
Finally and most importantly, I would like to thank God who by His grace gave me strength
and hope in all forms when I needed it most.
1
1. General introduction
1.1. Toxoplasma gondii
Toxoplasma gondii is a protozoan parasite that infects virtually all species of warm blooded
animals, including humans. The parasite can infect many different types of tissue, and can
cause symptoms ranging from subclinical infection to severe multi-systemic disease.
Infection with T. gondii is widely prevalent in humans and other animals on all continents
(Dubey, 2007).
The parasite was first named in 1908 when asexual stages were found in the tissues of a
laboratory rodent, Ctenodactylus gundii (Nicolle and Manceaux, 1908). The coccidian
nature of T. gondii was first exposed in the 1960s during electron microscopy studies which
revealed ultrastructural similarities between extraintestinal merozoites of T. gondii and
intestinal merozoites of Eimeria species (Levine, 1977). In the 1960s the heteroxenous
lifecycle of T. gondii was elucidated after it was found that the faeces of cats may contain an
infectious stage of T. gondii which induces infection when ingested by intermediate hosts
(Hutchison, 1965). Awareness of the coccidian life cycle of T. gondii was completed in the
1970s by the discovery of sexual stages in the small intestine of cats (Tenter et al., 2000).
After the initial discovery of T. gondii, several species of Toxoplasma were described,
mainly in accordance with the host species in which they were found (Levine, 1977). In
addition, several other protozoa were assigned to the genus Toxoplasma, but since then
these have been reclassified into other coccidian genera. During the past three decades, T.
gondii has been generally considered the only valid species of the genus Toxoplasma
(Tenter et al., 2000).
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1.2.Life cycle
1.2.1. Introduction
T. gondii is a coccidian with a heteroxenous life cycle. The parasite can infect a wide range
of tissue in warm blooded animals. T. gondii has a predilection for neural and muscle tissue
(Dubey et al., 1998), however can be widely dispersed in the body, particularly in acute
infection (Dubey and Beattie, 1988). There are three infectious stages of T. gondii; oocysts,
tachyzoites and bradyzoites (Figure 1.1). Tachyzoites multiply rapidly in tissues whereas
bradyzoites multiply slowly within tissue cysts. Both tachyzoites and bradyzoites are found
within host tissue. Oocysts are the environmental stage of T. gondii and only originate in the
faeces of felids.
Felids are the only definitive host of T. gondii and shed large numbers of oocysts briefly
during acute infection (Dubey and Frenkel, 1972). Oocysts are not immediately infective
and must first undergo sporulation in the environment, which may take 1 to 5 days (Dubey
et al., 1970). Other warm blooded animals, are capable of acting as intermediate hosts and
when infected may hold T. gondii bradyzoites and tachyzoites within their tissues. There are
three ways intermediate hosts and felids may become infected with T. gondii (i) the
ingestion of infective oocysts, (ii) the ingestion of viable bradyzoites present in tissue cysts
of an infected host and (iii) vertical (transplacental or transmammary) transmission of
tachyzoites from mother to offspring (Tenter et al., 2000).
3
Figure 1.1 Major routes of transmission of T. gondii (modified from (Tenter et al., 2000))
1.2.2. Oocyst transmission
In Australia, the introduced cat Felis catus, is the only known definitive host of T. gondii.
Felids shed oocysts in their faeces for only one to two weeks after initial infection with T.
gondii (Dubey and Frenkel, 1972, 1976). Although it is very rare for cats to re-shed oocysts
later in life, re-shedding has been reported in cats infected with other coccidian parasites,
following immunosuppression or after re-exposure to T. gondii years after initial infection
(Dubey, 1976, 1995; Dubey and Frenkel, 1974). Oocysts can remain infective in soil for up
to 2 years under favourable climatic conditions (Frenkel et al., 1975). Oocysts remain viable
4
for longer periods of time in a cool and moist environment (Yilmaz and Hopkins, 1972). As
most cats only shed once in their life, kittens and juvenile cats are the most common source
of T. gondii oocysts, as opposed to mature cats (Frenkel and Ruiz, 1981).
Hosts may ingest oocysts through the direct ingestion of cat faeces containing oocysts or the
ingestion of material contaminated with oocysts from cat faeces. Oocysts may be found in
water, soil and feed. Oocysts may also be transmitted mechanically by invertebrate
paratenic hosts. For example, earthworms are a source of T. gondii infection in eastern
barred bandicoots (Perameles gunnii) (Bettiol et al., 2000b; Obendorf and Munday, 1990).
Earthworms (Annelida) and beetles (Coleoptera) make up a significant proportion of the
diet of eastern barred bandicoots. It was proposed that a potential source of infection in
these animals is from ingestion of arthropods and earthworms which contain T. gondii
oocysts in their digestive tracts (Bettiol et al., 2000b; Obendorf and Munday, 1990). In an
experimental feeding study to assess the role of earthworms in the transmission of T. gondii
infection to eastern barred bandicoots, the findings confirmed that the earthworms
(Lumbricus rubellus and Perionyn excavatus) can transmit T. gondii infection (Bettiol et al.,
2000b). Oocysts present in the alimentary tracts of the worms, rather than infective stages of
T. gondii in worm tissues were responsible for the T. gondii infections in the bandicoots.
Marine bivalves such as mussels (Mytilus galloprovincialis) and oysters (Crassostrea
virginica) are also known to concentrate T. gondii oocysts (Arkush et al., 2003; Lindsay et
al., 2001) and were implicated as a source of T. gondii for Californian sea otters (Enhydra
lutris nereis) (Miller et al., 2008b).
5
1.2.3. Bradyzoite transmission
Animal tissue containing bradyzoites is another source of T. gondii infection. Tissue from a
T. gondii infected animal can contain bradyzoites in tissue cysts, which is infective when
eaten raw or undercooked (Dubey and Beattie, 1988). Bradyzoites in tissue cysts often
become non-viable after freezing at -20°C (Dubey, 1974). Tissue cysts may remain viable if
animal tissue is stored above -20°C and is uncooked (Jacobs et al., 1960). Epidemiological
studies demonstrate that meat ingestion is a risk factor for T. gondii infection in humans. For
example, a study of Seventh Day Adventists, who as a group follow a diet containing no
meat, found a significantly lower proportion of people in this group to be infected with T.
gondii compared to a control group (Roghmann et al., 1999). In addition, inhabitants of
France, particularly Parisians, who have a high consumption rate of rare meat, also have
among the highest rate of T. gondii infection in the world (Papoz et al., 1986).
1.2.4. Vertical transmission
Unlike oocyst and bradyzoite transmission, vertical transmission is generally not thought be
a major source of T. gondii infection in animal and human populations (Marshall et al.,
2004). Vertical transmission of T. gondii is traditionally thought to occur infrequently and
almost always in acutely infected pregnant females (Dubey and Beattie, 1988). The
terminology used to describe mother to young transmission of T. gondii varies. In this thesis
I will use the term ‘congenital transmission’ to describe the transmission of T. gondii from
dam to offspring in utero and the term ‘vertical transmission’ to refer to transmission
resulting from either transplacental or milk transmission (Miller et al., 2008a). Early studies
in mice and guinea pigs found that congenital infection with T. gondii can occur while the
dam is chronically infected with T. gondii (Remington et al., 1961). Despite this finding, it
6
is generally accepted in the literature that vertical transmission in all other animals is
infrequent and only occurs during acute infection. In humans, infection acquired before
pregnancy is thought to pose little or no risk to the foetus (Remington and Desmonts, 1990).
The immunological competence of the mother during parasitaemia and the number and
virulence of the parasites transmitted to the foetus is known to affect the risk of T. gondii
infection of the foetus and the severity of the disease (Tenter et al., 2000). The frequency of
transmission in humans varies according to time of gestation when the mother became
infected (Dunn et al., 1999). In addition, time of infection during gestation and severity of
disease are inversely related (Dunn et al., 1999). For instance, infection with T. gondii in the
first and second trimester more commonly results in severe congenital toxoplasmosis or
abortion. In contrast, late maternal infection in the third trimester usually results in
newborns without clinical signs of toxoplasmosis. The same dynamics of vertical
transmission of T. gondii in humans are thought to occur in sheep. Infection of sheep early
in gestation is rapidly fatal to the foetus due to the absence of the foetal immune response to
inhibit parasite multiplication (Buxton and Finlayson, 1986). Infection in mid-gestation may
also be fatal or give rise to a weak foetus. Infection late in pregnancy however will normally
cause foetal infection, but because at this stage the foetal sheep immune system is well
advanced, T. gondii will be resisted and the lamb is born infected and healthy (Buxton,
1990). Congenital transmission of T. gondii is reported in acutely infected cats (Dubey et
al., 1995a) and is reported in a number of other acutely infected animals including goats
(Dubey et al., 1985), pigs (Jungersen et al., 2001), dolphins (Jardine and Dubey, 2002) and
otters (Miller et al., 2008a). Recent studies in sheep have found that vertical transmission
occurs frequently (Duncanson et al., 2001; Williams et al., 2005) and it is increasingly being
proposed that vertical transmission may have significantly more influence on the prevalence
of T. gondii infections than was previously thought (Johnson, 1997).
7
Vertical transmission of T. gondii and its influence on the maintenance of T. gondii in
natural populations has been a matter of debate in recent years (Johnson, 1997). Vertical
transmission is commonly thought of in terms of its ability to cause abortion or debilitating
disease in the young, rather than its ability to contribute to the overall prevalence of T.
gondii infection (Dubey and Lappin, 1998; Duncanson et al., 2001; Marshall et al., 2004;
Tenter et al., 2000). Although it has long been known that that congenital infection with T.
gondii in mice and guinea pigs can occur while the dam is chronically infected (Remington
et al., 1961), recent studies have ignited debate as to whether vertical transmission is
common in other animals. Recent studies have verified the high frequency of congenital
transmission of T. gondii in chronically infected mice and it was proposed congenital
transmission in chronically infected mice can maintain T. gondii infection in wild mouse
populations (Marshall et al., 2004; Owen and Trees, 1998). In addition, a high frequency of
congenital T. gondii infection has recently been observed in naturally infected sheep in
which the resultant lambs were healthy (Duncanson et al., 2001). Recent data also suggests
T. gondii can be transmitted via successive vertical transmission within families of sheep
(Morley et al., 2005). Further studies need to be undertaken to determine the incidence of
vertical transmission in other chronically infected animals. If vertical transmission of T.
gondii does occur in several species of chronically infected animals and the resultant
offspring are healthy, this would suggest that vertical transmission is a more common
source of T. gondii infection that previously thought.
Unlike chronically infected mice, congenital toxoplasmosis in chronically infected rats is
extremely uncommon (Dubey et al., 1997b; Zenner et al., 1993). It is unknown what causes
the difference in result between different vertical transmission studies. A factor which is
8
known to affect the incidence of congenital T. gondii infection in humans and sheep is the
timing of infection in utero (Buxton, 1990; Dunn et al., 1999). In addition, the timing of
infection in utero is inversely related to the severity of disease in congenitally infected
humans and sheep (Buxton, 1990; Montoya and Liesenfeld, 2004). Additional factors in
animals that have been speculated to influence the probability and severity of congenital
infection with T. gondii include the size of the placenta, length of gestation,
immunocompetence of the foetus and maternal immunity (Johnson, 1997). Type II strains of
T. gondii tend to be found more commonly in cases of congenital T. gondii infection, and it
has been suggested that the genotype of T. gondii affects the likelihood and severity of
congenital infection (Darde et al., 2007). Further studies on the dynamics of vertical
transmission need to be undertaken to fully understand the role of vertical transmission in T.
gondii infection.
1.3.Pathogenesis and immunity
A number of factors may affect the pathogenesis of T. gondii in an intermediate host. These
include, inoculum size, genotype of T. gondii, immunological status and host species (Dardé
et al., 2008). When an intermediate host ingests tissue cysts or oocysts, the bradyzoites or
sporozoites are released into the lumen of the small intestine. These invade enterocytes or
intra-epithelial lymphocytes of the small intestine (Ferguson and Dubremetz, 2007). In
experimental infection of sheep, tachyzoites can be found multiplying in the mesenteric
lymph nodes by day 4 (Buxton et al., 2007). Tachyzoites are disseminated systemically via
the vascular system to most organs in the human body (Jackson and Hutchison, 1989).
Within an organ, tachyzoites infect host cells, multiply and invade adjoining cells.
9
Tachyzoite multiplication results in cell death and focal necrosis surrounded by an acute
inflammatory response (Bhopale, 2003).
Both a cellular and humoral immune response control T. gondii infection within the host.
Acute toxoplasmosis and associated multifocal necrosis can occur if tachyzoite
multiplication is not controlled by the host immune response, and this is known to occur in
human and ovine congenital toxoplasmosis and in mice infected with T. gondii of the type I
genotype (Dardé et al., 2008). In an immunocompetent host, tachyzoites stimulate
macrophages to produce IL-12, which in turn activates natural killer cells and T cells to
produce IFN-γ (Bhopale, 2003; Gazzinelli et al., 1993). IFN-γ and tumor necrosis factor
(TNF) act to mediate killing of tachyzoites by macrophages (Daubener et al., 1996; Sher et
al., 1993; Sibley et al., 1991). CD8+ T cells secrete IFN-γ and display in vitro cytotoxicity
towards T. gondii infected host cells (Khan et al., 1990; Subauste et al., 1991). CD4+ T cells
are also cytotoxic to T. gondii infected cells (Montoya et al., 1996) and produce IL-2 which
induces lymphokine activated killer cells which are also cytotoxic (Mosmann et al., 1986).
T helper 2 cells produce IL-4, IL-5 and IL-10 which are associated with down regulation of
cell mediated immune response (Mosmann and Moore, 1991). T. gondii also stimulates the
production of IgG, IgM, IgA and IgE antibodies against the parasite. Extracellular
tachyzoites are lysed by anti-T. gondii antibodies in the presence of complement (Sabin and
Feldman, 1948; Schreiber and Feldman, 1980). In addition, human platelets are cytotoxic to
tachyzoites (Yong et al., 1991). T. gondii seropositive animals are immune to T. gondii
infection, however infection persists as bradyzoites within tissue cysts (Buxton et al., 2007).
Reactivated toxoplasmosis may occur when a host is immunosuppressed, at which time
bradyzoites reconvert to tachyzoites and multiply.
10
Unlike the host immune response to tachyzoites, the host immune response to bradyzoites
within tissue cysts is minimal. The mechanisms that cause tachyzoite to bradyzoite
conversion are poorly understood (Bhopale, 2003; Dardé et al., 2008). Immune response
factors such as IFN-γ, TNF, IL-12 and T cells may indirectly control stage differentiation
(Dardé et al., 2008). Stage conversion has been examined in experimentally infected mice
(Ferguson and Dubremetz, 2007). The majority of tissue cysts are formed within striated
muscle and the central nervous system (Ferguson and Dubremetz, 2007). At 12-15 days
after oral infection, lesions are present in the brain which consist of tachyzoites, early tissue
cysts and inflammatory cells (Ferguson et al., 1991). The bradyzoites multiply within the
tissue cysts over three weeks and causes tissue cysts enlargement within the cell. Tissue
cysts are retained within a viable host cell. During chronic infection of mice, a very small
percentage of tissue cysts are found rupturing at any given time (Ferguson et al., 1989).
Tissue cyst rupture occurs during host cell death. In an immunocompetent host, cyst rupture
is associated with a large and rapid cell mediated immune response involving numerous
inflammatory cells (Ferguson and Dubremetz, 2007). In mice, macrophages have been
observed phagocytosing extracellular bradyzoites (Ferguson et al., 1989).
1.4. Diagnosis
Infection with T. gondii can be diagnosed in a number of ways. When tissue samples from
dead animals, whole blood, tissue biopsies or fluid aspirates are available, techniques such
as histology, immunohistochemistry, bioassays and polymerase chain reaction (PCR) can be
utilised to detect T. gondii organisms. When serum is available, serological techniques such
as the Sabin-Feldman dye test, modified agglutination test (MAT), enzyme-linked
immunosorbent assay (ELISA), indirect fluorescent antibody test (IFAT) and
11
immunoblotting can be used to detect T. gondii specific antibodies. Each technique has its
own costs and benefits and no single technique is 100% sensitive or specific.
Diagnosis using histology detects T. gondii organisms themselves and also detects
pathology associated with T. gondii infection. However, during chronic infection, T. gondii
is spread sparsely within tissues and is often difficult to detect with histology (Reddacliff et
al., 1993). Identification of tachyzoites is characteristic of active infection (Montoya and
Liesenfeld, 2004). In animals, T. gondii must be differentiated from other Apicomplexa
such as Neospora spp and Sarcocystis spp. Immunohistochemistry or PCR specific for T.
gondii would subsequently enable accurate identification of the tachyzoites visualised in
histological slides.
Immunohistochemistry refers to the identification of antigenic determinants of specific
substances (proteins) by the application of antibodies to histological sections (Canfield and
Hemsley, 2000). Immunohistochemistry can be used to differentiate T. gondii from other
Apicomplexa in histological slides where tachyzoites or bradyzoites are found.
Alternatively, immunohistochemistry may be used in histological slides where T. gondii
organisms may be difficult to find, such as in chronic T. gondii infection. The principle of
immunohistochemistry is similar to other serological techniques such as ELISA and IFAT.
A colour reaction is observed where T. gondii organisms are present in the histological
slides.
Another method used in the diagnosis of T. gondii infection is bioassay. This technique uses
a T. gondii naïve host to amplify viable (infective) T. gondii organisms present in the
patient’s tissue. T. gondii naive cats or mice are usually used as hosts (Dubey and Beattie,
12
1988). Tissue, often from a dead animal, is inoculated into the host and the host is then
tested for T. gondii infection. Mice which die post inoculation are tested for T. gondii, often
by impression smears of the brain and/or lung to identify T. gondii organisms. Alternatively,
cats are serologically tested for T. gondii antibodies post-inoculation and the faeces of
seropositive cats are subsequently examined for T. gondii oocysts. Bioassays are known to
be highly sensitive at detecting T. gondii infection (Hill et al., 2006) as large volumes of
tissue can be tested at one time and only a small amount of infective T. gondii is required to
infect a naïve cat or naive mice. Bioassays may also be used to amplify T. gondii organisms
present in a patient’s tissue prior to PCR identification and sequencing. Direct PCR without
the use of amplification methods such as bioassays may fail to detect T. gondii organisms
present in low numbers. PCR may be less sensitive than bioassay because only a small
amount of tissue can be tested using PCR and T. gondii organisms are often dispersed
sporadically throughout tissues. One disadvantage of bioassays is that they fail to detect
non-viable T. gondii in tissue. T. gondii organisms may become non-viable after tissue is
fixed in formalin or ethanol or after long periods or desiccation or freezing (Dubey and
Beattie, 1988). Other disadvantages are that bioassays require the use of laboratory animals
and are expensive and labour intensive.
PCR detection of T. gondii identifies T. gondii DNA present in tissue, however it can also
be used to identify T. gondii DNA present in whole blood, fluid aspirates and the faeces of
felids. PCR techniques for T. gondii have increased in popularity since the early 1990s
(Burg et al., 1989). Newer techniques which use nested primers and amplify high copy
DNA fragments within the T. gondii genome claim to be highly sensitive at detecting T.
gondii (Pujol-Rique et al., 1999). The main disadvantage of PCR detection of T. gondii is
that only a small amount of tissue can be tested at one time. T. gondii is often dispersed
13
sporadically within tissues, therefore a “needle in a haystack” scenario may ensue. A major
advantage of PCR is that once T. gondii DNA is detected, the genotype of T. gondii may be
identified via DNA sequencing and phylogenetic relationships can be made.
Another method of diagnosis of T. gondii infection is the detection of T. gondii specific
antibodies in serum. Serology is the method of diagnosis preferred in live animals as it can
detect antibodies during blood screening, which is far less invasive than tissue biopsy or
fluid aspiration. Serology is also relatively sensitive at detecting T. gondii infection
compared to bioassay, PCR and histology (Hill et al., 2006). The detection of IgM denotes
recent or active T. gondii infection whereas the detection of IgG implies chronic T. gondii
infection (Dardé et al., 2008). New serotyping techniques in humans have the ability to
diagnose the strain of T. gondii infecting a patient based on serology alone, and may be
applicable to animals in the future (Kong et al., 2003). A number of serological techniques
exist to detect T. gondii antibodies in serum, each with different sensitivities and
specificities. Serological techniques used to detect T. gondii infection in marsupials are
outlined in section 1.6.3.
Serology must be used with caution in neonates. This is because maternal antibodies
(transferred passively in utero or through milk) must be differentiated from the neonate’s
own antibodies. Serological testing of the neonate can be performed after the time when
maternal antibodies subside to help differentiate passive immunity from actual infection.
The time at which maternal antibodies subside is species dependent. Serological techniques
to differentiate maternal immunity from actual infection in neonates are used in humans
(Chumpitazi et al., 1995; Gavinet et al., 1997; Gross et al., 2000; Pinon et al., 2001;
Remington et al., 1985). Of these, comparative immunoblotting is the most popular
14
technique and the only technique used to date in cats (Cannizzo et al., 1996) for neonatal
serodiagnosis of T. gondii infection. Comparative immunoblots are theoretically applicable
to all species. Immunoblots detect T. gondii specific antibodies which bind to different
antigens ultimately causing a visible banding pattern on membrane strips. Comparative
immunoblots can therefore be used to compare the T. gondii immune response of the dam
with that of the offspring. If the banding pattern of the dam and offspring are the same, this
suggests maternal immunity is responsible for the offspring’s seropositivity. However, if the
offspring have independent bands from the dam, this would suggest the offspring was
producing its own antibodies against T. gondii and is actually infected.
1.5.Molecular epidemiology of T. gondii
1.5.1. Introduction
The majority of T. gondii isolates found to date have been grouped into three highly clonal
but closely related lineages, which differ in virulence and epidemiological pattern of
occurrence (Ajzenberg et al., 2004). A number of studies have linked the genotype of T.
gondii with a particular manifestation of infection. Congenital infection, ocular disease and
reactivated toxoplasmosis are three commonly described disease manifestations of T. gondii
infection in humans.
The terminology used in T. gondii molecular epidemiology is highly variable. In this thesis I
will use the terms ‘atypical’, ‘recombinant’ and ‘novel’ in the following way; ‘Atypical’
isolates fall into two general classes: ‘recombinant’ strains which have genotypes that are
clearly related to the three dominant types; and ‘novel’ strains which have a significant level
15
of polymorphism and often originate from wildlife or remote areas. In addition I will use the
term ‘isolate’ to refer to a sample from an individual animal or human, whereas I will use
the term ‘strain’ to refer to the genotype of T. gondii isolate, which can fall into ‘types’ I, II,
III or atypical.
1.5.2. Clonal lineages of T. gondii
A number of molecular studies of T. gondii isolates, predominately from domestic animals
and humans from Europe and North America, conclude that a majority of T. gondii isolates
comprise of three clonal lineages referred to as type I, II and III. Studies on T. gondii
lineages began with isoenzyme analysis and antigenic analysis and then progressed to using
molecular tools, particularly the polymerase chain reaction combined with restriction
fragment length polymorphism (PCR-RFLP), random amplified polymorphic DNA
polymerase chain reaction (RAPD-PCR), DNA sequencing and microsatellite DNA
analysis.
Studies have characterised T. gondii into mouse virulent and mouse avirulent lineages using
numerous loci and a variety of analyses. In an initial genetic study by Sibley and Boothroyd
(1992) PCR-RFLP analysis at the SAG-1, 850 and BS loci of 10 mouse virulent isolates
revealed an essentially identical genotype among the isolates and it was concluded that the
virulent isolates of T. gondii comprise a single clonal lineage. This genetically
homogeneous virulent lineage was also found on isoenzyme analysis which noted most
virulent isolates fell into a single zymodeme (Darde et al., 1992). RFLP analysis of DNA
polymerase α genes (Binas and Johnson, 1998) in addition to gene sequence data from
HSP70 (Lyons and Johnson, 1998) and reverse transcriptase PCR of SAG1 (Windeck and
16
Gross, 1996), all demonstrate the dichotomy of a virulent and avirulent lineage. In addition,
a study by Guo et al (1997) used RAPD-PCR to differentiate 35 T. gondii isolates into a
genotype of virulent strains and a genotype of avirulent strains. (Guo et al., 1997)
The existence of three clonal lineages was considered after a number of studies using multi-
locus PCR-RFLP. One of the first studies to suggest the existence of three clonal lineages
was Parmley et al (1994) which used RFLP analysis at three loci (P22, SAG1 and 850). The
result of virulent isolates being genetically identical and comprising a single lineage (group
A) was consistent with previous studies. However, heterogeneity seen among the 21
avirulent isolates was categorised into two genetically identical clonal lineages (group B and
C). A subsequent study by Howe and Sibley (1995) comprised a larger number of isolates
and produced similar findings. RFLP analysis was employed at six loci with 106 isolates. It
was found that virulent isolates were represented in one clonal lineage (type I) whereas
avirulent isolates were represented in two clonal lineages (type II and type III). It was
concluded by Howe and Sibley (1995) that T. gondii has a clonal population structure in that
>95% of isolates fall clearly into 1 of 3 distinct lineages. This theory of the highly clonal
population structure was further validated by studies using 8 microsatellite markers on 83
stocks (Ajzenberg et al., 2002a), sequencing of 7 single-copy genes on 16 stocks (Lehmann
et al., 2000) and sequencing of 15 loci on 18 stocks (Grigg et al., 2001a). However an
increasing number of isolates are being found, particularly in wildlife and geographically
isolated areas, that do not fit into the three distinct genotypes (Ajzenberg et al., 2004).
(Parmley et al., 1994)
17
1.5.3. The effect of T. gondii genotype on disease manifestation
It has been known for some time that certain strains of T. gondii (type I) are highly virulent
in mice, whereas others are avirulent. Isolates generally fit into two extremes in mice:
highly virulent, with an LD100 (the dose at which 100% of animals die) of one parasite, or
avirulent, with an LD100 of several thousand parasites (Boothroyd and Grigg, 2002). There
are however no guarantees that the differences in virulence seen in mice will also be seen in
other animals. Type I strains multiply approximately three times faster in human foreskin
fibroblasts than type II and III strains and this may give an indication that the differences
seen in mice may extend to humans and other animals (Boothroyd and Grigg, 2002). It is
unknown what strains are responsible for the bulk of human infections as most human
infections do not exhibit overt disease and form chronic cysts which cannot be genotyped by
testing bodily fluids. Tachyzoites present in severe disease are present at the site of disease,
and depending on the disease location can subsequently occupy the amniotic fluid, aqueous
humor or cerebrospinal fluid. It is therefore possible to access tachyzoites in severe
infections which subsequently enables molecular analysis and strain typing.
Type II strains of T. gondii tend to be found more commonly in cases of congenital T.
gondii infection in Europe and North America, and it has been suggested that the genotype
of T. gondii affects the likelihood and severity of congenital infection (Darde et al., 2007).
Analyses in France indicate that of 13 isolates collected from cases of congenital
toxoplasmosis all were type II strains (Howe et al., 1997). A limited analysis of samples
from the USA supported the trend of type II strains being most common in congenital
infection (Howe and Sibley, 1995). An additional study genotyped 86 samples, primarily
from France and Belgium using both mouse inoculation and microsatellite analysis
18
(Ajzenberg et al., 2002b). Isolates were characterised using eight microsatellite markers and
it was found that 85% were type II, 8% were type I, 3% were type III and 4% were atypical
genotypes. This study also analysed the relationship between T. gondii genotype and clinical
manifestations of patients with congenital toxoplasmosis. Type II isolates were predominant
among both the severe toxoplasmosis group as well as the group of patients with benign and
asymptomatic toxoplasmosis. In contrast, no type I T. gondii DNA was isolated from the
benign and asymptomatic group of patients (Ajzenberg et al., 2002b). Although the time the
foetus is initially infected with T. gondii plays a large role in the severity of infection in the
foetus, the data from this study suggests the strain of T. gondii may also play a role in
severity of disease. A conflicting study in Spain (Fuentes et al., 2001) identified type I T.
gondii was predominant in congenital toxoplasmosis. The bias in the Spanish study towards
sampling clinically severe cases of toxoplasmosis may explain this difference. In addition,
it is likely that there is a geographical variation in strains associated with congenital
toxoplasmosis. For example in Brazil, Colombia and French Guiana, the majority of T.
gondii DNA isolates characterised were type I, atypical or recombinant strains (Ajzenberg et
al., 2004; Ferreira et al., 2006). In the few reports of congenital T. gondii strain typing in
these countries, which were all from cases of severe congenital toxoplasmosis, recombinant
type I/III strains, type I or SAG1 type I strains were identified.
Type I T. gondii strains are strongly associated with ocular toxoplasmosis as shown by a
number of investigations in human patients. Data concerning ocular disease and its
association with type I strains are consistent and not conflicting. Vallochi et al (2005)
showed that parasite DNA isolated from all 11 ocular toxoplasmosis patients in Brazil were
from type I T. gondii strains. A study of USA patients (Grigg et al., 2001b) observed that in
rare occurrences of ocular toxoplasmosis in otherwise healthy adults, type I and/or
19
recombinant genotypes bearing the SAG1 type I allele (associated with mouse virulence)
were found in all six patients. Conversely, type II and III strains were only found to cause
ocular disease in immunosuppressed patients. Similar results were seen in Canada (Burnett
et al., 1998) and Brazil (Glasner et al., 1992) where isolates from ocular toxoplasmosis
outbreaks were found to be type I strains (Boothroyd and Grigg, 2002).
Toxoplasmosis in Brazil seems to differ from other countries in that a majority of isolates
originating from Brazil have been genotyped as type I, recombinants of type I or novel
strains. This is in contrast to studies performed in the US and Europe in which most isolates
were avirulent types II or III. The fact that a majority of Brazilian isolates genotyped are
closely related to the type I lineage may be important finding (Ferreira et al., 2006). From a
recent study of 20 Brazilian isolates, 85% showed a significant degree of virulence (Ferreira
et al., 2006). Speculation arose that the high frequency of type I isolates found in Brazil may
be in part responsible for the high frequency of acquired ocular toxoplasmosis in humans in
Brazil (Ferreira et al., 2006; Khan et al., 2006). Cases of ocular toxoplasmosis in Brazil are
often recurrent and serious in nature (Glasner et al., 1992; Silveira et al., 2001).
The relationship between T. gondii genotype and severity of infection was also investigated
in sea otters in the Californian coast. Infection with T. gondii and associated
meningoencephalitis was recognised as a major cause of death in subadults and prime-aged
adult sea otters, accounting for 16% of total otter mortality (Miller et al., 2004). A novel
type X strain was identified to predominate in all infected otters, being present in 72% of all
beach-cast otters examined by genotypic analysis (Conrad et al., 2005). It was found that
type X infected otters tended to have moderate to severe meningoencephalitis on
histopathology more frequently than type II infected otters (Miller et al., 2004). In addition,
20
more otters infected with type X T. gondii had T. gondii associated meningoencephalitis as a
primary cause of death when compared with type II infected otters (Miller et al., 2004).
1.5.4. Atypical T. gondii genotypes
It is widely thought that T. gondii has a low genetic diversity due to the common finding of
strains in Europe and North America that can be grouped into three highly clonal but closely
related lineages (Howe and Sibley, 1995; Johnson, 1997; Su et al., 2003). However, it is
increasingly being proposed that the genetic diversity among T. gondii isolates worldwide is
greater than current estimates. One reason may be sampling bias that has resulted from the
study of isolates from humans and domestic animals primarily originating from North
America and Europe.
Recombinant strains of T. gondii have alleles identical to those found in the three major
lineages but these alleles have segregated differently among the loci analysed (Darde et al.,
2007). Recombinant genotypes are related to the three main lineages. Isolates with mixed
genotypes were sampled from areas including Brazil (Ferreira et al., 2006), Africa, the
Caribbean and Reunion Island (Ajzenberg et al., 2004). A few recombinant strains to date
were isolated from wildlife, and these were from bears and deer in North America
(Ajzenberg et al., 2004; Howe and Sibley, 1995). Although the finding of clonal strains is
by far the most common in T. gondii isolated from humans and domestic animals in North
America and Europe, some recombinants are found. These recombinants include four
isolates from North American pigs (Mondragon et al., 1998a), five samples from human
ocular toxoplasmosis patients in North America (Grigg et al., 2001b), two samples from
AIDS patients in the USA (Howe and Sibley, 1995) and four isolates from human
21
congenital toxoplasmosis patients in France (Ajzenberg et al., 2004; Ajzenberg et al.,
2002b).
Novel strains of T. gondii have many unique polymorphisms and novel alleles (Darde et al.,
2007). Just fourteen novel strains are described in the literature to date. The first novel strain
found was MAS, isolated from a case of human congenital toxoplasmosis in France and the
second found was CASTELLS isolated from an aborted sheep in Uruguay (Darde et al.,
2007). The other atypical strains cited in the literature are a cougar isolate from Canada
(Lehmann et al., 2000), type X from marine mammals in the USA (Conrad et al., 2005;
Miller et al., 2004), isolate IPP-2002-URB from a human congenital toxoplasmosis patient
in France (Ajzenberg et al., 2004) and nine strains from French Guiana (Ajzenberg et al.,
2004).
A majority of the novel or recombinant strains isolated to date were found from areas
outside Europe and North America or in non-domesticated animals. In contrast, many of the
T. gondii isolates that were used to propose the majority of T. gondii isolates fall into three
clonal lineages have been collected from human patients and domestic animals in Europe
and North America (Ajzenberg et al., 2004; Lehmann et al., 2006). In light of this
knowledge it is possible that the genetic diversity of T. gondii is highly underestimated. The
collection of isolates tested for the clonal theory may not reflect the true status of T. gondii
in previously unsampled regions such as in remote geographical areas.
A number of studies which have genotyped T. gondii isolates from wildlife (Dubey et al.,
2004a) and from geographically isolated locations (Dubey et al., 2002; Dubey et al., 2003a;
Dubey et al., 2003b; Dubey et al., 2003c; Dubey et al., 2003d) have not identified novel
22
genotypes. Many of these studies have used one PCR-RFLP marker to identify isolates as
either type I, II or III. Studies which have identified novel or recombinant isolates of T.
gondii (Bossi et al., 1998; Carme et al., 2002; Darde et al., 1998; Howe and Sibley, 1995;
Lehmann et al., 2000; Miller et al., 2004) have used techniques including isoenzyme
analysis, microsatellite analysis, multilocus PCR-RFLP and gene sequencing.
Misidentification of unusual recombinants or novel isolates can commonly result from the
use of a single genetic marker in genotype analysis. For example, in the molecular
characterisation of T. gondii DNA isolated from ocular toxoplasmosis patients, RFLP
analysis at any one locus would have misidentified the 5 recombinant isolates found (Grigg
et al., 2001b). In addition, analysis at three loci (SAG1, SAG2 and SAG4) would have
misidentified an isolate (2035) as type I when it was another recombinant (Grigg et al.,
2001b). Therefore, it is likely that the many studies to date which have genotyped T. gondii
isolates using a small number of genetic markers have misidentified the strain of T. gondii
or have missed novel isolates.
Several studies have shown an unusual T. gondii population structure in Brazil. PCR-RFLP
at eight independent loci was used to determine the clonal lineage of 20 T. gondii isolates
from humans and animals in Brazil (Ferreira et al., 2006). The finding that 100% of T.
gondii isolates analysed from this population were natural recombinants was different from
the expected frequency. Previous studies have reported that regardless of the host and
geographical origin, approximately 95% of T. gondii isolated belong to one of three
genetically distinct lineages (Darde et al., 1992; Howe and Sibley, 1995). Several studies
which used single locus PCR-RFLP of Brazilian T. gondii strains have reported a high
frequency of types I and III and an absence of type II (de A. dos Santos et al., 2005; Dubey
et al., 2003a; Dubey et al., 2003b; Dubey et al., 2004). The study by Ferreira et al (2006)
23
illustrates the usefulness of multilocus PCR-RFLP in identifying hybrid strains of T. gondii.
It was concluded by Ferreira et al (2006) that even the analysis of two loci may lead to the
misidentification of the genotype of Brazilian isolates. For example, if all Brazilian isolates
in the study were analysed using the SAG1 and B1 loci, they would have been identified as
being the type I lineage.
Studies to date have shown only restricted deductions can be made from individual
polymorphic markers. Detection of recombination events and interpretation of T. gondii
population structure often requires multilocus genotyping and the sensitivity of analysis
increases with the number of markers used (Darde et al., 2007). Sensitive and efficient
RFLP assays often used in older studies of T. gondii population genetics assume T. gondii is
composed of only three clonal lineages. These assays may misclassify isolates representing
new lineages and certain recombinants. The large number of novel and recombinant isolates
found in wildlife and in areas outside of North America and Europe suggest that the genetic
diversity of T. gondii is higher than previously estimated (Howe and Sibley, 1995). Testing
of T. gondii isolates from wildlife and isolated areas using multilocus genotyping may
reveal a large proportion of recombinant and novel isolates. No studies to date have
described the molecular characterisation of T. gondii isolates from wildlife in Australia and
it is of special interest considering Australia’s isolation and unusual wildlife species.
1.6.Significance of T. gondii in Australian marsupials
Australian marsupials are among the most susceptible hosts for T. gondii and the parasite is
known to cause both chronic and acute infections (Basso et al., 2007; Beveridge, 1993). It is
commonly thought that marsupials are highly susceptible to manifesting disease when
24
exposed to T. gondii due to their lack of evolutionary exposure to felids (Innes, 1997). There
are no native felids present in Australia, and cats were only introduced during European
settlement. Infection in marsupials is not always fatal and can result in long-term latent
infection which can be reactivated during times of stress (Obendorf and Munday, 1983).
Tissue cysts present in latent infection can reactivate at a later time and cause clinical
disease which normally presents as neurological deficits (Lynch et al., 1993b). T. gondii
infection may make a marsupial more prone to predation by affecting its movement,
coordination and sight (Obendorf and Munday, 1983, 1990). Toxoplasmosis is associated
with widespread pathology and death in several collections of captive marsupials (Barrows,
2006; Boorman et al., 1977; Canfield et al., 1990; Dobos-Kovacs et al., 1974; Dubey et al.,
1988; Hartley, 2006; Hartley et al., 1990; Miller et al., 1992; Patton et al., 1986). Captivity
is a stressor and may therefore increase the chance of reactivated T. gondii infection
(Arundel et al., 1977; Beveridge, 1993; Obendorf and Munday, 1983, 1990).
1.6.1. Life cycle of T. gondii in Australian marsupials
Marsupials may become infected with T. gondii through feed contaminated with oocysts or
via the ingestion of tissues containing bradyzoites. Oocyst contamination of stored feed
stuffs and food containers are often blamed for toxoplasmosis outbreaks in captive
marsupials (Dubey et al., 1988; Miller et al., 1992; Patton et al., 1986). However, it is also
possible that captive marsupials can become infected with T. gondii and remain healthy long
term until a stressor causes an outbreak of reactivated toxoplasmosis (Obendorf and
Munday, 1990). As mentioned above, omnivorous marsupials such as eastern barred
bandicoots can become infected with T. gondii via ingestion of contaminated earthworms
containing oocysts in their digestive tracts (Bettiol et al., 2000b). Carnivorous marsupials
25
may become infected with T. gondii via ingestion of T. gondii-infected predated animals or
bradyzoites in raw meat. Carnivorous marsupials include species in the family Dasyuridae;
such as chuditch (Dasyurus geoffroii), kowaris (Dasyuroides byrnie), brush-tailed
phascogales (Phascogale tapoatafa) and Tasmanian devils (Sarcophilus harrisii). In
captivity, dasyurids fed on diets of fresh meat are prone to T. gondii infection (Attwood et
al., 1975). Infection with T. gondii in kangaroos can also become a public health issue as
kangaroo meat is consumed by humans and domestic pets (Holds et al., 2008). Kangaroo
meat is commonly enjoyed rare and kangaroo pet meat is regularly served raw. As T. gondii
bradyzoites remain infective when meat is undercooked, the ingestion of rare or raw
kangaroo meat is a risk factor in T. gondii transmission (Robson et al., 1995). T. gondii-
infected kangaroo meat is not only a source of infection for humans, but also for domestic
cats, which may subsequently shed oocysts and perpetuate the life cycle.
The incidence of vertical transmission in marsupials is yet to be determined; however it is of
special interest considering the impact of toxoplasmosis in marsupials. Evidence for vertical
transmission in marsupials to date is anecdotal (Boorman et al., 1977; Dubey et al., 1988).
Dubey et al. (1988) describes two black-faced kangaroo (Macropus fuliginosus melanops)
dams with positive MAT results and T. gondii-infected pouch young. Both pouch young
died, one at 82 days of age and the other at 7 months of age, and toxoplasmosis was
confirmed in both using histology. It is highly unlikely that the pouch young tested in this
study were exposed to T. gondii oocysts from the external environment as both died before
first pouch exit (Dawson, 1995). Marsupial young first exit the pouch after a long period of
permanent residence and, while within the pouch, are protected from the external
environment (Tyndale-Biscoe and Renfree, 1987). Congenital transmission was also
suspected in an outbreak of toxoplasmosis in wallaroos (Macropus robustus) (Boorman et
26
al., 1977). Of four wallaroos that died of toxoplasmosis, three were 6 months of age. The
other wallaroo that died of toxoplasmosis was the dam of one of the dead pouch young. Of
the three pouch young with toxoplasmosis, one was hand reared from 5 months of age and
had the potential to be infected with T. gondii from the external environment. The other two
pouch young were not hand reared and were unlikely to be infected with T. gondii from the
external environment as they were 6 months old and the approximate age of first pouch
young exit in wallaroos is 7 months.
T. gondii transmission via the milk is the most likely mechanism of vertical transmission in
marsupials as opposed to transplacental infection. Milk transmission is likely because
marsupial young are born at a very immature state (less than 1gram neonatal weight)
(Tyndale-Biscoe and Renfree, 1987) and milk is the source of nourishment which enables
the young to develop to a state where they can leave the pouch (Dawson, 1995). Therefore if
pouch young are infected in utero, they are not likely to survive initial infection (Dubey et
al., 1988). Vertical transmission via the milk has recently been proposed to play a role in the
life cycle of T. gondii (Johnson, 1997). Milk transmission of T. gondii is not well
documented, however tachyzoites have been isolated from the milk of a number of species
including mice, cats, cows, pigs, dogs, sheep, rats, guinea pigs and rabbits (Johnson, 1997).
Tachyzoites are infectious orally to cats and mice (Dubey, 1998) which suggests that
tachyzoites in milk are infectious via the gastrointestinal route. In addition, in experimental
infections of lactating mice acid-resistant T. gondii bradyzoites were found in the milk and
were able to produce consistent infection via the gastrointestinal route (Pettersen, 1984).
Several studies have suggested congenital transmission via the milk is common in cats
(Dubey, 1995; Powell et al., 2001; Powell and Lappin, 2001). The transmission of T. gondii
in humans through breastfeeding was suspected in a mother who suffered clinical signs of
27
acute toxoplasmosis after pregnancy and whose suckling child was subsequently
seropositive for anti-T. gondii IgM (Bonametti et al., 1997). Humans may also become
infected with T. gondii by drinking unpasteurised goat’s milk (Riemann et al., 1975; Sacks
et al., 1982).
1.6.2. T. gondii associated disease in Australian marsupials
Although T. gondii infection is commonly implicated as a cause of death in captive
marsupials, the impact of T. gondii infection in wild marsupials is more difficult to
determine as predation of recently infected marsupials hinders investigation into the cause
of death. In-depth investigations regarding the impact of T. gondii in eastern-barred
bandicoots led to the conclusion that T. gondii infection is a significant cause of death
among this species, both in captivity and in the wild (Bettiol et al., 2000a; Miller et al.,
2000; Obendorf and Munday, 1990; Obendorf et al., 1996). In 1984, reports of a CNS
disease affecting eastern barred bandicoots were received from two locations in Tasmania.
Several bandicoots were observed with signs of incoordination, apparent blindness,
unnatural daytime activity and erratic staggering movements (Obendorf and Munday, 1990).
These bandicoots eventually died and necropsy results confirmed toxoplasmosis as the
cause of death in several wild bandicoots (Obendorf and Munday, 1990; Obendorf et al.,
1996). In addition, T. gondii was speculated to cause deaths in the wild in the common
brushtail possum (Trichosurus vulpecula) (Eymann et al., 2006) and Tasmanian pademelon
(Thylogale billardierii) (Obendorf and Munday, 1983). A case report of toxoplasmosis in
wild Tasmanian pademelons, where two carcasses were examined histologically, found T.
gondii to be the cause of death (Obendorf and Munday, 1983). These two wallabies were
found stumbling blindly and were subsequently euthanased. According to the land owner,
28
sick and dead wallabies had been observed every year, with the number of wallabies
affected increasing yearly (Obendorf and Munday, 1983).
Clinical signs of toxoplasmosis in Australian marsupials vary and include diarrhoea,
respiratory distress, weight loss, blindness, neurological deficits and sudden death (Miller et
al., 2003). Several species of marsupial have been found to be infected with T. gondii using
histology (Ashton, 1979; Attwood et al., 1975; Barrows, 2006; Basso et al., 2007; Canfield
et al., 1990; Dubey et al., 1988; Hartley, 2006; Hartley et al., 1990; Miller et al., 1992;
Obendorf and Munday, 1983; Obendorf et al., 1996; Patton et al., 1986; Skerratt et al.,
1997). The histopathology of T. gondii infection is highly variable and can range from no
identifiable lesions to severe multisystemic necrosis. Descriptions of pathological lesions of
T. gondii infection in marsupials are incomplete and limited. Detailed descriptions of lesions
in dasyurids (Attwood et al., 1975), koala (Phascolarctos cinereus) (Hartley et al., 1990),
sugar glider (Petaurus breviceps) (Barrows, 2006), common wombat (Vombatus ursinus)
(Hartley, 2006; Skerratt et al., 1997), eastern barred bandicoot (Bettiol et al., 2000a) and a
number of macropod species (Basso et al., 2007; Canfield et al., 1990; Dubey et al., 1988;
Miller et al., 1992; Obendorf and Munday, 1983; Patton et al., 1986; Reddacliff et al., 1993)
have been published. A review of the pathology of 79 naturally infected marsupials,
including macropods, common wombats, koalas, possums, dasyurids, numbats
(Myrmecobius fasciatus), bandicoots and a bilby (Macrotis lagotis) was also published
(Canfield et al., 1990). In this review lungs were commonly affected with congestion and
oedema or interstitial pneumonia and macrophage accumulation. Myocardial, skeletal and
smooth muscle necrosis and neutrophilic inflammation were common. The adrenals,
pancreas and liver often showed focal areas of necrosis and fibrinous exudate. In addition,
the tissues of the central nervous system (CNS) commonly showed focal necrosis. The
29
stomach and small intestine showed mucosal ulceration and often extensive smooth muscle
necrosis. T. gondii tissue cysts were common in muscle and nervous tissue and free
tachyzoites were common in areas of necrosis. Similar pathological changes were observed
in a study by Obendorf and Munday (1983) which describes acute toxoplasmosis in
naturally infected wild macropods.
A study of the pathology of experimentally induced toxoplasmosis in macropods
(Reddacliff et al., 1993) described slightly different pathological changes. Nine tammar
wallabies (Macropus eugenii) were infected with T. gondii oocysts orally. Seven of the 9
died acutely of toxoplasmosis whereas two survived with chronic toxoplasmosis. In all
wallabies with acute toxoplasmosis, prominent histological changes were seen in the small
intestine, lungs and mesenteric lymph nodes. The most extensive areas of necrosis and
largest numbers of tachyzoites were seen in the gastrointestinal tract and mesenteric lymph
nodes. Necrotic lesions in other organs, including the CNS, were much less extensive and
tissue cysts were not detected. Tachyzoites were mostly seen in areas of necrosis or
inflammation. In the two chronically infected tammar wallabies, minimal histological
lesions were observed apart from occasional small foci of inflammation in the brain, heart,
skeletal muscle and liver. No tissue cysts were observed using histology in these
chronically infected animals despite careful searching of serial sections. T. gondii infection
of the two chronically infected animals was confirmed by mouse bioassay. Similar
pathology was observed in a colony of zoo macropods with serological evidence of acute T.
gondii infection (Patton et al., 1986).
Publications describing toxoplasmosis in marsupials demonstrate that T. gondii associated
pathology is extremely variable. Nevertheless, common findings in cases of acute
30
toxoplasmosis in marsupials are focal areas of necrosis and/or inflammation in areas such as
the muscle, viscera and/or CNS. T. gondii organisms were more common in areas of
necrosis and inflammation, but there were exceptions. Ocular toxoplasmosis is reported in
wallabies (Ashton, 1979). In some cases it is necessary to perform PCR or bioassay from
tissues which are suspected of infection with T. gondii but where no organisms are seen.
PCR or bioassay detection of T. gondii in tissue is particularly important in animals which
are chronically infected with T. gondii, which may not have significant pathology.
Additionally PCR or immunohistochemistry may be used to differentiate T. gondii infection
from infections with other Apicomplexa such as Sarcocystis species and Neospora species.
1.6.3. Diagnosis of T. gondii infection in Australian marsupials
Immunohistochemistry is successfully used to diagnose T. gondii infection in a number of
marsupial species (Barrows, 2006; Basso et al., 2007; Canfield et al., 1990; Hartley, 2006;
Hartley et al., 1990). Bioassays are also successfully used to detect T. gondii in marsupials
(Basso et al., 2007; Johnson et al., 1989; Reddacliff et al., 1993). A limited number of T.
gondii serological techniques are applied to marsupials. The MAT is the most commonly
used test for T. gondii specific IgG antibodies in Australian marsupials (Dubey et al., 1988;
Hartley and English, 2005; Lynch et al., 1993b; Miller et al., 2003; Miller et al., 2000) and
is the only test routinely used to screen marsupials for T. gondii infection in zoos throughout
Australia. Published studies show a good correlation between MAT positivity in marsupials
and infection with T. gondii (Johnson et al., 1989; Obendorf et al., 1996). The popularity of
the MAT in marsupials stems from the test not utilizing a secondary reagent to detect T.
gondii antibodies, so enabling it to be used on a range of marsupial species. In addition, the
MAT is used extensively for the diagnosis of toxoplasmosis in a range of other species
31
(Dubey, 2007) and is used as a sensitive and specific test to detect T. gondii IgG antibodies
in humans (Desmonts and Remington, 1980), mice (Dubey et al., 1995b), pigs (Dubey et al.,
1995c), sheep (Ljungstrom et al., 1994) and felids (Dubey et al., 1995a; Dubey et al.,
2004b). The MAT is available as a commercial kit (Toxo-Screen DA, bioMerieux, Marcy
l’Etoile, France). An ELISA was developed to detect anti-T. gondii IgG in marsupials
(Johnson et al., 1988) however it was not made available commercially and is not
commonly used, with no further publications mentioning its use. Compared to agglutination
tests, which can test a large range of species, ELISAs and IFATs can often only be applied
to one species at a time due to their use of species-specific reagents. No commercial ELISA
or IFAT is available for use in any species of marsupial, however reagents are available if
one chooses to create an in-house ELISA or IFAT. An advantage of the ELISA is that its is
high throughput and results can be easily interpreted based on the cut off point for optical
density (Johnson et al., 1988).
The only published method describing the detection T. gondii-specific IgM in marsupials to
date is the direct agglutination test (DAT) (Johnson et al., 1989). The DAT is similar to the
MAT, however in the MAT, 2-mercaptoethanol (2-ME) is added to destroy non-specific
antibodies and IgM (Desmonts and Remington, 1980). In theory, the difference in titre
between the MAT and DAT will demonstrate if T. gondii specific IgM antibodies are
present in sera. Johnson et al (1989) demonstrated the use of the DAT in the serodiagnosis
of acute toxoplasmosis in macropods. Samples from 17 Tasmanian pademelons and 17
Bennett’s wallabies (Macropus rufogriseus rufogriseus) were used to correlate the presence
and absence of T. gondii in the brain (via bioassay) with DAT results. In addition, three
eastern grey kangaroos (Macropus giganteus) were experimentally infected with T. gondii
and their serological response recorded via the DAT and MAT. Results showed that the use
32
of the DAT and MAT on marsupials may diagnose acute toxoplasmosis. Many studies have
since used the DAT and MAT to detect IgM in marsupials (Bettiol et al., 2000a; Bettiol et
al., 2000b; Hartley and English, 2005; Hartley, 2006; Lynch et al., 1993a; Lynch et al.,
1993b; Miller et al., 2000; Obendorf et al., 1996; Skerratt et al., 1997). Although the DAT is
not available commercially it can be easily made by omitting the addition of 2-ME in the
commercially available MAT.
Other serological tests that are used to detect anti-T. gondii antibodies in marsupials are the
Sabin-Feldman dye test (Dubey et al., 1988) and the latex agglutination test (Dubey et al.,
1988; Hartley and English, 2005; Turni and Smales, 2001). The Sabin-Feldman dye test is
the gold standard for the serodiagnosis of T. gondii in humans (Reiter-Owona et al., 1999),
but has had limited use in marsupials. Although the dye test produced similar results to the
MAT in detecting anti-T. gondii IgG in macropods (Dubey et al., 1988), its complexity,
need for special reagents and use of live infective parasites are the likely reasons for it’s
unpopularity in marsupial T. gondii serodiagnosis. The latex agglutination test (LAT) has
also had limited use in marsupials and has a lower sensitivity to detect IgG in macropods
compared to the MAT (Dubey et al., 1988). The latex agglutination test has varying
sensitivity and specificity in different studies and species examined (Dubey et al., 1985;
Dubey et al., 1988; Mazumder et al., 1988). Of all serological tests used in marsupials, cut
off values are only established for the DAT (Johnson et al., 1989) and an in-house ELISA
(Johnson et al., 1988). The cut off point titre for the DAT (1:64) is commonly imposed upon
MAT results (Hartley and English, 2005; Hartley, 2006; Miller et al., 2000; Obendorf et al.,
1996), however a cut off point of 1:25 is also used for the MAT (Eymann et al., 2006) and
the cut off point recommended in the protocol for the commercially available MAT is 1:40.
33
Serodiagnosis of T. gondii infection in neonates can be difficult due to the presence of
maternal antibodies in some species. In marsupials, maternal anti-T. gondii antibodies must
be differentiated from the pouch young’s own antibodies. The same is true in human
neonates. A study by Yadav (1971) discovered maternal antibodies subside by the end of
pouch life in the quokka (Setonix brachyurus) and possum (Trichosurus vulpecula). The
time of permanent pouch exit in marsupials is species specific. However, the time of
permanent pouch exit is always slightly before the time of weaning. Maternal antibodies are
primarily transferred via the milk in marsupials (Old and Deane, 2000) and it is therefore
expected that maternal antibodies subside close to the time of weaning. Comparative
immunoblots were not used in published studies to detect T. gondii infection in young
marsupials, however are used in humans and cats (Cannizzo et al., 1996) and may be
applicable to all species.
1.6.4. Prevalence of T. gondii in Australian marsupials
A limited number of seroprevalence studies have been undertaken in wild Australian
marsupial populations. T. gondii seroprevalence in free ranging marsupials was 3.3% in
Bennett’s wallabies and 17.7% in Tasmanian pademelons using an ELISA (Johnson et al.,
1988), and 15% in bridled nailtail wallabies (Onychogalea fraenata) using a latex
agglutination test (Turni and Smales, 2001). In addition, T. gondii seroprevalence levels of
6.7% in eastern barred bandicoots (Obendorf et al., 1996), 26.1% in common wombats
(Hartley and English, 2005) and 6.3% in the common brushtail possum (Eymann et al.,
2006) were observed using the MAT. Seroprevalence results therefore indicate that some
wild marsupials are infected with T. gondii and survive.
34
Outbreaks of toxoplasmosis are of particular importance for rare and endangered
marsupials, especially those in captive breeding programs and in small free ranging
populations in remnant habitats (Lynch et al., 1993b). Further knowledge of the prevalence
and transmission of T. gondii in marsupials is warranted to better understand the dynamics
of infection in marsupials and to further develop management strategies to control
toxoplasmosis. Australian marsupials are well known to characteristically exhibit T. gondii-
related disease, however no previous studies to date have attempted to identify the genotype
of T. gondii that infect marsupials in Australia. Recombinant and novel isolates of T. gondii
are commonly associated with unusual clinical manifestations (Ferreira et al., 2006; Miller
et al., 2004). In addition, previous studies illustrated that isolated areas of the world tend to
harbour higher rates of atypical T. gondii genotypes. A study of the molecular epidemiology
of T. gondii in Australian marsupials may identify new, novel or recombinant strains of T.
gondii and could later assist in investigations that link different strains of T. gondii with
different disease manifestations.
1.7.Aims of this thesis
Infection with T. gondii is an important cause of disease and death in Australian marsupials.
However, little is known about the prevalence, transmission and strains of T. gondii in wild
Australian marsupials. Not only is the prevalence of T. gondii in wild marsupials of
importance in terms of conservation, the presence of infection in wild kangaroos in
particular is of public health significance due to the kangaroo meat trade (Holds et al.,
2008).
35
While it is plausible that environmental contamination with oocysts from cats is the sole
source of T. gondii in populations of herbivorous marsupials, it is also possible that vertical
transmission plays a role in the maintenance of T. gondii infection in marsupials. Evidence
for vertical transmission in marsupials to date is anecdotal (Boorman et al., 1977; Dubey et
al., 1988), and the incidence of vertical transmission in marsupials is unknown. Information
on the frequency of vertical transmission in marsupials will benefit captive breeding
programmes by ensuring that only T. gondii-free animals are bred, thereby improving
animal health and assisting animal conservation.
Knowing which strain(s) of T. gondii infects wild marsupials in Australia is also of
importance to wildlife conservation and management. Different strains of T. gondii differ in
virulence and are linked to different disease manifestations in humans and animals. No
studies to date are published that molecularly characterise T. gondii from wild Australian
marsupials. Therefore the aims of this study were to:
1. Develop a cost effective ELISA to detect T. gondii IgG in macropods;
2. Identify the seroprevalence of T. gondii in a range of wild marsupial species and
populations;
3. Evaluate the occurrence of vertical transmission of T. gondii in Australian
marsupials;
4. Determine the molecular characteristics of T. gondii DNA found in wild Australian
marsupials.
36
1.8.Study design
An initial seroprevalence study was organised to assess the prevalence of anti-T. gondii IgG
in wild Australian marsupials. An ELISA was developed to detect anti-T. gondii IgG in
macropod serum. Sera was collected from a range of marsupial species in different locations
nation wide and tested for anti-T. gondii IgG using an ELISA and the MAT. Sera was
collected in order to assess the differences in T. gondii seroprevalence between different
marsupial species and to ascertain the effect of location on seroprevalence. In addition to a
general seroprevalence study, sera and/or tissue samples were obtained from marsupial
dams and their corresponding pouch young. Paired dam-pouch young samples were
obtained in order to determine the frequency of vertical transmission of T. gondii in
marsupial species. The type of sample obtained was dependent on the state of the animal
when sampled. In marsupials where samples were obtained within hours of death, namely
culled western grey kangaroos (Macropus fuliginosus), both sera and tissues were sampled
from dams and their young in pouch. In captive marsupials that were alive when sampled,
only sera was obtained from both dam and pouch young. For welfare reasons, live captive
young were only bled after the time of natural pouch exit. In marsupials that were found
dead with young in pouch, only tissue samples were obtained. Serum samples from dam-
pouch young pairs were tested using a number of techniques. Firstly, paired sera were
screened for T. gondii IgG using the MAT or ELISA. Comparative immunoblots were then
utilised to attempt to differentiate passive immunity from actual infection in seropositive
pouch young. In addition, seropositive dam sera were tested using both the MAT and DAT
to determine the presence of IgM. Tissue samples were preserved for both PCR and
histology. PCR was utilised as a diagnostic test in dams and their pouch young whereas
37
histology was used to detect T. gondii related pathology. PCR products were also sequenced
in order to analyse the molecular characteristics of the T. gondii DNA found.
38
2. The development of an in-house ELISA for the detection of
anti-T. gondii antibodies in macropod marsupials
2.1.Introduction
Infection with T. gondii can be diagnosed in a number of ways. Diagnosis using histology
and bioassays detect T. gondii organisms themselves, but require tissue from dead animals.
Furthermore, during chronic infection, T. gondii is spread sparsely within tissues and is
often difficult to detect with histology (Reddacliff et al., 1993). Bioassays, although highly
sensitive at detecting T. gondii infection, are expensive and labour intensive (Hill et al.,
2006). PCR detection of T. gondii DNA also necessitates invasive sampling techniques or
necropsy. Serology identifies serum antibodies which are easy to detect during routine blood
screening. The presence of anti-T. gondii IgG in sera is indicative of chronic T. gondii
infection in marsupials (Hartley, 2006; Johnson et al., 1989).
During studies which involved the screening of western grey kangaroos (Macropus
fuliginosus) for antibodies against T. gondii, a commercially available modified
agglutination test (MAT) (Toxo-Screen DA, bioMerieux, France) was used. The MAT was
chosen to screen initial serum samples because it is the most commonly used test for
serodiagnosis of T. gondii infection in Australian marsupials (Dubey et al., 1988; Hartley
and English, 2005; Lynch et al., 1993b; Miller et al., 2003; Miller et al., 2000) and is the
only test routinely used to screen marsupials for T. gondii infection in zoos throughout
Australia. Published studies show a good correlation between MAT positivity in marsupials
and infection with T. gondii (Johnson et al., 1989; Obendorf et al., 1996). The popularity of
the MAT in marsupials stems from the test not utilizing a secondary reagent to detect anti-T.
39
gondii antibodies, thus enabling it to be used on a range of marsupial species. In addition,
the MAT is used extensively for the diagnosis of toxoplasmosis in a range of other species
and is used as a sensitive and specific test to detect anti-T. gondii IgG antibodies in humans
(Desmonts and Remington, 1980), mice (Dubey et al., 1995b), pigs (Dubey et al., 1995c),
sheep (Ljungstrom et al., 1994) and cats (Dubey et al., 1995a; Dubey et al., 2004b). During
routine screening of macropod species for T. gondii antibodies, the MAT was found to be
cost prohibitive. Therefore, a cost effective in-house ELISA (enzyme-linked immunosorbent
assay) which detects anti-T. gondii IgG in macropod marsupials was developed. This
ELISA was found to be in high agreement with the MAT. Absolute agreement was
subsequently found between ELISA and T. gondii PCR results of western grey kangaroos
2.2.Materials and methods
2.2.1. Sample collection
Sera were obtained from three species of macropod to optimise and validate the ELISA.
Forty five sera samples from agile wallabies (Macropus agilis) and twelve sera samples
from eastern grey kangaroos (Macropus giganteus) were provided to Murdoch University
by staff at Rockhampton Zoo, QLD. Once obtained, serum samples were stored at -20oC.
Western grey kangaroo (Macropus fuliginosus) blood samples were obtained from an initial
54 kangaroos (group A) culled during Department of Environment and Conservation (DEC)
population control programmes in Perth, WA. Kangaroos were culled in areas such as parks,
reserves, golf courses and farms due to overpopulation. Blood was collected by needle
aspiration of the heart within 4 hours of death of the kangaroo and stored a 4oC for a
40
maximum of 2 days prior to centrifugation. Sera was separated from the blood clot and
stored at -20oC.
Paired blood and tissue samples were then collected from an additional 62 western grey
kangaroo dams and their offspring (group B) during the kangaroo culling programmes in
Perth, WA. An ID was allocated to each animal and blood, brain and tongue samples were
collected. Blood was collected by needle aspiration of the heart within 4 hours of death of
the kangaroo. Sera was separated by centrifugation and stored at -20oC. The head of each
adult kangaroo was removed in the field and transported to the laboratory and stored at 4°C
for a maximum of 3 days prior to processing. Samples of brain and tongue were then
removed from the head of adult kangaroos, placed in sterile containers and frozen at -20°C.
The pouch young of all 62 kangaroos were also killed in line with DEC population control
measures, via blunt trauma to the head. Samples of brain and heart were removed from the
pouch young once transported to our laboratory. Tissue samples were placed in sterile
containers and frozen at -20°C. All sera samples were tested using the ELISA.
2.2.2. Modified agglutination test
Fifty four serum samples from western grey kangaroos (group A), twelve serum samples
from eastern grey kangaroos and forty five agile wallabies were tested using the
commercially available MAT (Toxo-Screen DA, bioMerieux, France). Sera were tested at
two different sera dilutions; 1:40 and 1:4000, according to the manufacturer’s instructions.
The positive and negative control sera included in the kit were used in each round of
samples tested, in addition to an antigen control comprised of PBS (phosphate buffered
saline), according to the manufacturer’s protocol. A serum sample was determined to be T.
41
gondii positive when an agglutination reaction was observed at a serum dilution of at least
1:40.
2.2.3. Cell culture of T. gondii tachyzoites
Antigen for the ELISA was prepared from RH strain T. gondii tachyzoites grown in Vero
cell culture. All reagents and instruments used in tissue culture were sterile and all
procedures were carried out aseptically in a tissue culture cabinet. Reagents used in tissue
culture were heated to 37°C before use. Monolayers of Vero cells were grown in 25cm2 cell
culture flasks (Corning Incorporated, Corning, USA). Growth medium for the Vero cell
culture was DMEM (Dulbecco’s Modified Eagle’s Medium AQmedia, Sigma-Aldrich,
Castle Hill, Australia) plus 10% foetal bovine serum (DKSH, Hallam, Australia), 2mM L-
glutamine (Sigma-Aldrich, Castle Hill, Australia), 50ug/ml streptomycin and 50 IU/ml of
penicillin (Sigma-Aldrich, Castle Hill, Australia). Maintenance medium was identical to
growth medium, except the concentration of foetal bovine serum was lowered to 2%. When
a monolayer of Vero cells was confluent, all media was removed from the flask with a
pipette and 10ml PBS added. The cells were rinsed in PBS and the PBS then removed with
a pipette after which 1ml of trypsin was added to the flask and incubated at 37°C for a
maximum of 10 minutes. When the Vero cells were dislodged from the flask wall, 6ml of
sterile growth media was added and mixed with the Vero cells by gentle pipette action.
Using a pipette, 1ml of the resulting Vero cell suspension was added to 4 new flasks, each
of which contained 6ml of growth media. The flasks were stored in a humidified incubator
at 37°C, 5% CO2 for 2 days until a confluent monolayer was reached. Growth media was
then removed with a sterile pipette and 6ml of maintenance media added. Of the flasks, one
was kept as a stock of clean Vero cells and three were inoculated with T. gondii tachyzoites.
42
Cell culture flasks containing a confluent monolayer of Vero cells were inoculated with
approximately 5x 104 T. gondii fresh RH strain tachyzoites each. After 3-4 days, Vero cells
were lysed sufficiently and the tachyzoites harvested and purified. A cell scraper was used
to scrape Vero cells containing tachyzoites from the flask wall. The resulting suspensions of
T. gondii infected Vero cells in maintenance media were removed with a pipette and pooled.
Tachyzoites were purified from the Vero cells by shearing through a 30G needle then
filtration with a 5μm syringe filter (Pall Corporation, East Hills, USA). Purified tachyzoites
were washed twice in PBS pH 7.2. The suspension of tachyzoites in PBS was then sonicated
for 3 periods of 1 minute at a power level of 5 (SonicatorR Ultrasonic Processor, Misonix
incorporated, Farmingdale, USA). A Bradford protein assay (Quick Start TM Bradford Dye
Reagent, Biorad Laboratories, Gladesville, Australia) was undertaken on a pooled amount
of sonicated antigen and the antigen concentration adjusted with a volume of PBS to
produce 1000μg/ml of protein.
2.2.4. ELISA development
MAT tested sera were used to optimise the ELISA. The optimum concentrations of antigen,
serum and reagents for the ELISA were determined using a checker board system with
antigen diluted in one direction and a series of different sera and reagent concentrations
diluted in opposite directions. Four dilutions of antigen, 1μg/ml, 10μg/ml, 25μg/ml,
50μg/ml, were tested, with two MAT positive and two MAT negative sera samples diluted
at 1:100, 1:200 and 1:400 (Figure 2.1). The dilution of serum and antigen with the highest
difference between MAT positive and MAT negative sera samples were then selected for
use in assays to determine the optimum concentration of secondary and tertiary reagents.
43
Three dilutions of secondary reagent, 1:500, 1:1000 and 1:2000, were tested against three
dilutions of tertiary reagent, which were 1:1000, 1:2000 and 1:4000 and a serum dilution of
1:400 was used with an antigen concentration of 1 μg /ml (Figure 2.2). Two MAT positive
and two MAT negative western grey kangaroo sera were used in the initial optimisation
assays. After the optimal antigen and reagent concentrations were identified, 1 positive and
5 negative sera samples were tested at 8 sequential dilutions from 1:400 to 1:51200, using a
secondary and tertiary reagent dilution of 1:1000 (Figure 2.3). The serum dilution with the
greatest difference between positive and negative was chosen as the serum dilution for use
in the ELISA; this was a serum dilution of 1:800. After the concentration of antigen,
reagents and sera were optimised, a total of 111 MAT tested macropod serum samples were
used to validate the ELISA and establish a positive cut-off point for optical density (OD).
The cut-off point was determined by calculating the mean optical density plus 2 standard
deviations (SD) of MAT negative sera, this was 0.636. A serum sample with an OD reading
equal to the mean plus 2 SDs of MAT negative sera was used as a cut-off positive control.
For control of plate to plate variation, 2 negative and 2 positive control sera, including the
cut-off positive control, were included on every plate. A serum sample was considered to be
positive when its OD value was greater than that of the cut-off positive control, as described
in Stanley et al, (2004). (Stanley et al., 2004)
The final protocol for the ELISA commenced with T. gondii antigen diluted in 50mM
carbonate buffer, pH 9.6, to a concentration of 1 ug/ml. One hundred microliters of diluted
antigen was then added to every well of a 96 well ELISA plate (Microlon 600, Greiner Bio-
one, Germany). The 96 well plate was incubated for 1 hour at 37oC. A washing cycle
followed which consisted of rinsing in PBS with 0.05% Tween 20 for three periods of 3
minutes. The ELISA plate was blocked for 1 hour using 150 ul of 5% skim milk powder in
44
PBS 0.05% Tween 20 and washed. Duplicates of test sera samples diluted in 5% skim milk
in PBS were added at a volume of 100 ul and concentration of 1:800. Two MAT
seropositive and two MAT seronegative sera samples were included in every 96 well plate
tested. Sera was incubated for 90 minutes at 37oC and the plate washed prior to the addition
of 100ul of commercially available unconjugated rabbit anti-kangaroo IgG (Kangaroo IgG
(h&I) Antiserum, Bethyl Laboratories Inc, Montgomery, USA) at a concentration of 1:1000.
The ELISA plate was then incubated for a further 60 minutes and washed, after which 100
ul of Horseradish Peroxidase (HRP)-conjugated anti-Rabbit antibody (Donkey Anti-Rabbit:
HRP, Affinity BioreagentsTM, Golden, USA) was added at a concentration of 1:1000. After
the final incubation period of 60 minutes, the plate was washed and 200 ul OPD (o-
phenylenediamine Dihydrochloride) Substrate Solution (Sigma FastTM OPD, Sigma-
Aldrich, Castle Hill, Australia) added. The OPD was left in the dark at room temperature for
15 minutes before the reaction was stopped with 50 ul of 2M H2SO4. The optical density
was then read at 450nm using a spectrophotometer.
2.2.5. ELISA validation
Serum samples tested using the MAT that were used to optimize the ELISA were retested
using the final ELISA protocol. In addition, serum samples collected from western grey
kangaroos in group B were screened using the final ELISA protocol. PCR specific for T.
gondii DNA was then used to test tissue samples from ELISA positive and ELISA negative
western grey kangaroos in group B.
45
2.2.6. DNA extraction
DNA from tissue of 9 ELISA positive and 9 ELISA negative western grey kangaroos from
group B was extracted for polymerase chain reaction (PCR). Frozen tissue samples were
thawed and then homogenised using sterile instruments and containers. A number of
methods of DNA extraction were used. The MasterPure DNA purification kit (Epicentre
Biotechnologies, Madison, USA) was used initially as it was more cost effective to use
compared to QIAGEN kits (QIAGEN, Hilden, Germany). An in-house DNA extraction
protocol was later used for proceeding samples obtained to further increase cost
effectiveness. The QIAamp DNA MiniKit (QIAGEN, Hilden, Germany) and
phenol/chloroform extraction was then used to extract DNA from tissue samples. This was
because although QIAamp DNA MiniKit and phenol/chloroform extraction are more
expensive than the previously used methods, they are thought to better reduce PCR
inhibitors than other methods (Dean et al., 2004; Pinto et al., 2007).
DNA samples were extracted using the MasterPure DNA purification kit according to the
manufacturer’s directions, 5mg of tissue was used in each extraction. In the in-house DNA
extraction protocol 100mg of each sample was incubated overnight at 37°C in 2ml cell lysis
buffer (0.1M Tris-HCl2, 0.01M ethylene diamine tetra-acetic acid, 1% sodium dodecyl
sulfate, pH 8) containing proteinase K (Sigma-Aldrich, Castle Hill, Australia) at a final
concentration of 150ug/ml. After overnight incubation, 60ug RNase A (Sigma-Aldrich,
Castle Hill, Australia) was added to the suspension and the samples incubated for a further
30 minutes at 37°C. Samples were then placed on ice for a minimum of 3 minutes and 1ml
of 7.5M ammonium acetate added. Each sample was then vortexed for 10 seconds and
pelleted by centrifugation for 10 minutes at 2000 x g. The resulting supernatant was
46
transferred to a clean tube and 3ml isopropanol added. The tube was inverted 40 times and
the DNA pelleted by centrifugation for 10 minutes at 2000 x g. The DNA pellet was washed
in 70% ethanol, dried and resuspended in 30ul TE buffer solution pH 7.4 (Fisher Biotec,
Wembley, Australia) and stored at -20°C.
A method of phenol-chloroform DNA extraction was also used to extract DNA. Briefly,
25mg of homogenised tissue was incubated at 37°C for 2 hours in 300ul of cell lysis buffer
containing proteinase K (Sigma-Aldrich, Castle Hill, Australia). The DNA in the tissue
suspension was extracted using phenol/chloroform (1:1). DNA was precipitated with 30ul of
3M sodium acetate in 825ul 100% ethanol. Following washing in 70% ethanol, the DNA
pellet was resuspended in 30ul TE buffer (Fisher Biotec, Wembley, Australia). The QIAamp
DNA MiniKit was used according to the manufacturer’s directions. Water was used as a
negative control in the DNA extractions and RH strain T. gondii in Vero cells were used as
a positive DNA extraction control in each round of DNA extraction.
2.2.7. PCR
Extracted DNA was tested using PCR. Since the sensitivity of the PCR depends on the copy
number of the gene amplified (Switaj et al., 2005), the ITS1 sequence (110 copies) and the
B1 gene (35 copies) which are both present in high copy numbers, were chosen for use.
Both the ITS1 sequence and B1 gene have sequences that are specific for T. gondii. A
nested PCR for the B1 gene (Grigg and Boothroyd, 2001) was used after it was found that
many tissue samples from T. gondii seropositive animals were PCR negative using non-
nested primers (Bretagne et al., 1993). DNA from the RH strain of T. gondii was used as a
positive PCR control and PCR negative controls consisted of distilled water. Neospora
47
caninum DNA was tested using PCR primers for the B1 gene (Bretagne et al., 1993; Grigg
and Boothroyd, 2001) prior to testing sample DNA in order to confirm the specificity of the
primers used. Each sample of DNA was tested twice using each set of primers.
The nested PCR primers for the ITS1 sequence used (Nandra and Grigg, manuscript in
preparation) have the potential to amplify DNA from a range of Apicomplexa including T.
gondii gondii, Sarcocystis neurona, Neospora caninum, Hammondia hammondi and
Besnoitia species. Primers were designed so that PCR products of T. gondii were 440 base
pairs in size whereas PCR products of other Apicomplexa were of a different size. PCR
reactions were performed using 25ul volumes with the final mix containing 1ul template
DNA, 10 pMol of each primer, 0.2mM dNTPs, 2.5ul PCR buffer (Taq DNA Polymerase
10x Reaction Buffer, Fisher Biotec, Wembley, Australia), 3.75mM MgCl2, 0.6 units of Taq
Polymerase (Tth Plus* DNA Polymerase, Fisher Biotec, Wembley, Australia).
Amplification consisted of denaturing at 94°C for 5 minutes followed by 35 cycles of 94°C
for 40 seconds, 58°C for 40 seconds and 72°C for 90 seconds, after which there was an
extension period of 10 minutes at 72°C. PCR products were visualized using 0.8% agarose
gels stained with ethidium bromide. A 100bp DNA ladder (Promega, Madison, USA) was
included in each agarose gel. PCR products that were approximately 440 base pairs in size
were sequenced to identify a T. gondii specific DNA sequence.
One PCR assay used to amplify the T. gondii B1 gene used non-nested primers (Bretagne et
al., 1993). In the optimised protocol, 25ul reaction volumes were used with 1ul of template
DNA, 12.5 pMol of each primer, 0.2mM dNTPs, 2.5ul PCR buffer (Taq DNA Polymerase
10x Reaction Buffer, Fisher Biotec, Australia), 3.75mM MgCl2, 0.5 units of Taq
Polymerase (Tth Plus* DNA Polymerase, Fisher Biotec, Australia). Amplification consisted
48
of denaturing at 95°C for 5 minutes followed by 40 cycles of 94°C for 30 seconds, 65°C for
30 seconds and 72°C for 60 seconds, after which there was an extension period of 10
minutes at 72°C. PCR products were visualized using 0.8% agarose gels stained with
ethidium bromide. A 100bp DNA ladder (GeneRuler 100bp DNA ladder, Fermentas,
Burlington, Canada) was included in each agarose gel.
An additional nested PCR for the T. gondii B1 gene was used (Grigg and Boothroyd, 2001).
Briefly, 1ul of template DNA was added to a total reaction volume of 25ul, which consisted
of 10 pMol of each primer, 0.2mM dNTPs, 2.5ul PCR buffer (Taq DNA Polymerase 10x
Reaction Buffer, Fisher Biotec, Australia), 3.75mM MgCl2, 0.6 units of Taq Polymerase
(Tth Plus* DNA Polymerase, Fisher Biotec, Australia). Amplification consisted of
denaturing at 95°C for 5 minutes followed by 30 cycles of 94°C for 40 seconds, 60°C for 40
seconds and 72°C for 90 seconds, after which there was an extension period of 10 minutes
at 72°C. PCR products were visualized using 0.8% agarose gels stained with ethidium
bromide. A 100bp DNA ladder (Promega, Madison, USA) was included in each agarose gel.
PCR products were cut from agarose gels and DNA was purified from agarose using the
UltraClean GelSpin DNA Extraction Kit (MO BIO Laboratories Inc, Carlsbad, USA)
according to the manufacturer’s directions. Sequencing reactions were performed using
a BigDye Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems, Scoresby, Australia)
according to the manufacturer’s directions and using internal primers PCR primers.
Reactions were electrophoresed through an ABI 3730 automatic sequencer and sequencing
profiles analysed using FinchTV version 1.4 (Geospiza, Seattle, USA).
49
2.2.8. Statistics
Agreement between the MAT and ELISA and the ELISA and PCR was estimated by κ
coefficient (Smith, 1995).
2.3.Results
The in-house ELISA was in very high agreement with the MAT as illustrated in Table 2.1,
and yielded a kappa value of 0.96. Out of 111 kangaroo and wallaby sera samples tested,
only 2 discordant results were obtained, both of which were from haemolysed sera from
agile wallabies that were positive on the MAT but negative on the ELISA (Table 2.1).
Twenty two out of 24 MAT positive agile wallaby serum samples were positive on the
ELISA and all 21 MAT negative agile wallaby serum samples were negative on the ELISA.
Complete agreement was observed in seven MAT positive and 47 MAT negative western
grey kangaroo sera samples (Group A) that were tested on the ELISA. In addition, complete
agreement between the MAT and ELISA was observed in eastern grey kangaroos, with 2
MAT positive eastern grey kangaroo serum samples being positive on the ELISA and 10
MAT negative eastern grey serum samples being negative on the ELISA (Table 2.1).
Neospora caninum DNA was not amplified using primers for the B1 gene (Bretagne et al.,
1993; Grigg and Boothroyd, 2001). The results of the PCR of western grey kangaroo (group
B) tissue were in absolute agreement with the ELISA results (Table 2.2). T. gondii specific
DNA was detected in all nine western grey kagaroos that had sera which was ELISA
positive. In addition, all tissue samples from ELISA negative kangaroos were PCR negative
for T. gondii DNA using primer sets for both the B1 gene (Bretagne et al., 1993; Grigg and
50
Boothroyd, 2001) and ITS1 gene (Nandra and Grigg, manuscript in preparation). All PCR
products were sequenced, and BlastN analysis of the DNA sequences revealed that the
amplicons were specific for T. gondii.
2.4.Discussion
One hundred and eleven MAT-tested serum samples were used to validate and optimize the
ELISA developed in this study. This ELISA had a comparable sensitivity and specificity to
the MAT, based on its Kappa value of 0.96. A limited number of comparative studies are
published comparing the different methods used for serodiagnosis of T. gondii infection in
marsupials. One study utilized four different serological tests to test sera from seven black-
faced kangaroos and found that the MAT and the Sabin-Feldman dye test were more
sensitive at detecting T. gondii antibodies in kangaroo sera than both the indirect
agglutination and the latex agglutination tests (Dubey et al., 1988). The Sabin-Feldman dye
test is the gold standard for the serodiagnosis of T. gondii in humans (Reiter-Owona et al.,
1999), but has limited use in marsupials. Although the dye test was equivalent to the MAT
in detecting T. gondii IgG in macropods (Dubey et al., 1988), its complexity, need for
special reagents and use of live infective parasites are the likely reasons for it’s unpopularity
in marsupial T. gondii serodiagnosis. The principle advantage of the ELISA is that it can be
used to screen large numbers of serum samples more cost effectively than the commercially
available MAT. Another advantage of this ELISA is that its results can be easily interpreted
based on the cut off point for optical density, as compared to the IFAT (indirect fluorescent
antibody test) where slides need to be examined by experienced readers. Due to the
relatively high serum dilution of 1:800 used in this ELISA protocol compared to the MAT
serum dilution of 1:40, only a small amount of marsupial serum is required to detect T.
51
gondii antibodies using the ELISA. This is of particular benefit in wildlife research as only
small volumes of sera are usually obtainable and these are often used to test for multiple
conditions.
Rabbit anti-kangaroo IgG was used as the secondary reagent for the ELISA and the ELISA
worked not only for kangaroos but also for agile wallabies. It is unknown if the ELISA
developed is applicable to other marsupials, particularly those not within the genus
Macropus. Only one other paper has been published concerning the use of an ELISA to
detect T. gondii antibodies in macropods (Johnson et al., 1988). Sera from 17 Tasmanian
pademelons (Thylogale billardierii) and 17 Bennett’s wallabies (Macropus rufogriseus
rufogriseus), the brains of which had been bioassayed for T. gondii, were used to validate
the ELISA. Sheep anti-kangaroo immunoglobulin was used as a secondary reagent and it
was found that the ELISA was successful in detecting T. gondii antibodies in both Thylogale
billardierii and Macropus rufogriseus rufogriseus.
Absolute agreement between T. gondii ELISA results and the ITS1 PCR results was
observed in the 18 western grey kangaroos tested using both ELISA and PCR. Other studies
found PCR to be less sensitive than serology at detecting T. gondii infection in pigs (Garcia
et al., 2008; Hill et al., 2006) and cattle (More et al., 2008). The PCR used in this study
detecting T. gondii DNA in all 9 ELISA positive western grey kangaroos, which could
indicate that marsupials possess higher tissue burdens of parasites than for instance, pigs
and cattle. All ITS1 PCR products with correct sized bands for T. gondii were sequenced,
which subsequently confirmed T. gondii DNA was present in the tissue samples of
seropositive animals. T. gondii DNA was not detected in the tissues tested of 9 seronegative
kangaroos, which consisted of 3 adults and 6 pouch young. Although T. gondii can be
52
detected in seronegative animals (Owen and Trees, 1998), this is reported infrequently. PCR
results from DNA extracted from seropositive and seronegative kangaroos correlated
exactly with the serology results suggesting the ELISA developed is sensitive and specific.
Table 2.1Level of agreement between a commercially available MAT and an ELISA in western greykangaroos, eastern grey kangaroos and agile wallabies
Western grey kangaroo
(Group A)
ELISA
MAT + -
+ 7 0
- 0 47
Eastern grey kangaroo ELISA
MAT + -
+ 2 0
- 0 10
Agile wallaby ELISA
MAT + -
+ 22 2
- 0 21
MIXED TOTAL ELISA
MAT + -
+ 31 2
- 0 78
Kappa= 0.96
53
Table 2.2PCR results of ELISA positive and negative western grey kangaroos (Group B)
PCR results
Animal ELISA result Brain Tongue Heart
Adult C14 Positive B1, ITS1 Negative nd
Adult C9 Positive B1, ITS1 ITS1 nd
Adult J6 Positive B1, ITS1 ITS1 nd
Adult J10 Positive nd B1, ITS1 nd
Adult R7 Positive B1, ITS1 Negative nd
Adult Q1 Positive Negative B1, ITS1 nd
Adult G21 Positive B1 ITS1 nd
Adult F19 Positive ITS1 Negative nd
Adult R19 Positive ITS1 Negative nd
Adult F8 Negative Negative Negative nd
Adult H14 Negative Negative Negative nd
Adult I14 Negative Negative Negative nd
Pouch young 15B1 Negative Negative nd Negative
Pouch young R4 Negative Negative nd Negative
Pouch young F8 Negative Negative nd Negative
Pouch young H14 Negative Negative nd Negative
Pouch young I14 Negative Negative nd Negative
Pouch young Q20 Negative Negative nd Negative
B1- Positive B1 PCR (Bretagne et al., 1993)B1- Positive B1 PCR (Grigg and Boothroyd, 2001)ITS1- Positive ITS1 PCR (Nandra and Grigg, manuscript in preparation)Negative- Negative on all PCRs, nd- No sample available
54
Figure 2.1. Checker board system to determine initial serum and antigen dilutions for an ELISA
Abbreviations:Pos1- MAT positive serum sample 1Pos 2- MAT positive serum sample 2Neg1- MAT negative serum sample 1Neg 2- MAT negative serum sample 2
55
Figure 2.2. Checker board system to determine secondary and tertiary dilutions for an ELISA
Abbreviations:Pos1- MAT positive serum sample 1Pos 2- MAT positive serum sample 2Neg1- MAT negative serum sample 1Neg 2- MAT negative serum sample 2PBS- phosphate buffered saline, pH7.6
56
Figure 2.3. Checker board system to determine final serum dilution for an ELISA
Abbreviations:Pos1- MAT positive serum sample 1Neg1- MAT negative serum sample 1Neg 2- MAT negative serum sample 2Neg 3- MAT negative serum sample 3Neg 4- MAT negative serum sample 4Neg 5- MAT negative serum sample 5
57
3. Seroprevalence of T. gondii in free ranging Australian
marsupials
3.1.Introduction
Australian marsupials are among the most susceptible hosts for T. gondii and the parasite is
known to cause both chronic and acute infection (Basso et al., 2007; Beveridge, 1993).
Infection in marsupials is not always fatal and can result in long-term latent infection which
can be reactivated during times of stress (Beveridge, 1993; Obendorf and Munday, 1983). T.
gondii infection may make a marsupial more prone to predation by affecting its movement,
coordination and sight (Dubey and Beattie, 1988; Gonzalez et al., 2007). Not only is
infection with T. gondii attributed to causing declines in marsupial populations in the wild
(Eymann et al., 2006; Obendorf et al., 1996), toxoplasmosis is associated with widespread
pathology and death in several collections of captive marsupials (Barrows, 2006; Boorman
et al., 1977; Canfield et al., 1990; Dobos-Kovacs et al., 1974; Dubey et al., 1988; Hartley,
2006; Hartley et al., 1990; Miller et al., 1992; Patton et al., 1986). Captivity is a stressor and
therefore increases the chance of reactivated T. gondii infection (Arundel et al., 1977;
Beveridge, 1993; Obendorf and Munday, 1983). Clinical signs of toxoplasmosis in
Australian marsupials vary and include diarrhoea, respiratory distress, weight loss,
blindness, neurological deficits and sudden death (Miller et al., 2003). Common
histopathological findings include interstitial pneumonia of the lungs and myocardial,
skeletal and smooth muscle necrosis, with T. gondii cysts and tachyzoites in areas of
necrosis (Canfield et al., 1990). Due to the dynamics of T. gondii infection in marsupials,
knowledge of the T. gondii serological status of marsupials is of immense benefit to their
58
management in captivity and in the wild. For example, seropositive animals should be
managed in a way so as to reduce stressors in order to reduce the chance of reactivated
toxoplasmosis. A stressor that may induce reactivated toxoplasmosis in seropositive
marsupials includes capture stress (Obendorf and Munday, 1983).
Although a number of cases of toxoplasmosis are described in captive marsupials, there are
few recent data on the prevalence and distribution of T. gondii infection in wild marsupials.
T. gondii seroprevalence in free ranging marsupials was 3.3% in Bennett’s wallabies
(Macropus rufogriseus rufogriseus) and 17.7% in Tasmanian pademelons (Thylogale
billardierii) using an ELISA (Johnson et al., 1988), and 15% in bridled nailtail wallabies
(Onychogalea fraenata) using a latex agglutination test (Turni and Smales, 2001). In
addition, T. gondii seroprevalence levels of 6.7% in eastern barred bandicoots (Perameles
gunnii) (Obendorf et al., 1996), 26.1% in common wombats (Vombatus ursinus) (Hartley
and English, 2005) and 6.3% in the common brushtail possum (Trichosurus vulpecula)
(Eymann et al., 2006) were observed using the MAT. Not only is the prevalence of T. gondii
in wild marsupials of importance in terms of conservation, the presence of infection in wild
kangaroos in particular is of public health significance due to the kangaroo meat trade.
Kangaroo meat sourced from wild kangaroos is sold for human consumption in Australia,
Asia, Europe and North America (Holds et al., 2008).
In this study the prevalence of anti-T. gondii IgG was determined in a range of marsupial
species from a number of different locations. Western grey kangaroos (Macropus
fuliginosus) from Perth, WA and eastern grey kangaroos (Macropus giganteus) from
Sydney, NSW and Roma, QLD were tested using an ELISA. In addition, serum collected
from several populations of woylies (Bettongia penicillata) from throughout Western
59
Australia and South Australia was tested using the MAT. A seroprevalence study was also
undertaken in populations of marsupials and native rodents located in Faure Island, WA and
Barrow Island, WA. In addition meat eating wild chuditch (Dasyurus geoffroii) were tested
for anti-T. gondii IgG using the MAT.
The screening of wild kangaroos for exposure to T. gondii is important from a public health
perspective as kangaroo meat is consumed by humans and domestic felids (Holds et al.,
2008; Robson et al., 1995). An outbreak of toxoplasmosis was linked to the ingestion of rare
kangaroo meat served at a cocktail party and involved 12 acute infections in adults and a
case of congenital toxoplasmosis (Robson et al., 1995). T. gondii bradyzoites are more
likely to remain infective when meat is undercooked, making the ingestion of rare or raw
meat a risk factor in T. gondii transmission. T. gondii-infected kangaroo meat is not only a
source of infection for humans, but also to domestic cats, which can subsequently shed
oocysts and perpetuate the life cycle.
In addition to screening wild kangaroos, T. gondii seroprevalence was determined in woylie
populations. Woylies, like kangaroos are herbivorous macropods, but are classed in a
different subfamily to kangaroos and wallabies. The woylie is in the subfamily Potoroinae
whereas kangaroos and wallabies are both in the subfamily Macropodinae (Lee and
Cockburn, 1985). Woylies were sampled in locations where cats are free to roam in addition
to areas where cats are not located, namely St Peter Island, SA and Venus Bay Island, SA.
There is no history of cats being present on Venus Bay Island or St Peter Island (Van
Weenen, personal communication, November 19, 2007).
60
Populations of wild marsupials and rodents in Faure Island Sanctuary and Barrow Island
Nature Reserve were tested for anti-T. gondii IgG as part of this study. Animals exist on
these islands free from the presence of felids. Faure Island has been free from felids since
June 2001 (Thomas and Whisson, 2002) whereas there is no history of cats being present on
Barrow Island (Butler, 1982). In addition, the majority of land-dwelling mammals on these
islands were herbivorous. The only known way T. gondii could be maintained in
herbivorous populations without the presence of felids is via vertical transmission. It is
unknown if T. gondii infection can be maintained in populations of herbivorous animals free
from cats. In this study we determined the seroprevalence of T. gondii in cat free marsupial
populations in order to ascertain if T. gondii can be maintained in herbivorous populations
without the presence of felids.
A number of wild chuditch were tested for anti-T. gondii IgG. Chuditch are a meat eating
marsupial. It is thought that meat eating marsupials have a higher prevalence of T. gondii
than non-meat eating marsupials due to the presence of T. gondii bradyzoites in infected
meat (Obendorf and Munday, 1990). A high T. gondii prevalence of 51% was recorded in
wild and captive carnivorous dasyurid marsupials (Attwood et al., 1975). Wild chuditch
have also been found to have a moderate seroprevalence of T. gondii in the past with 14
chuditch out of 69 (20.3%) being seropositive in 1993 (Haigh, 1994). In this study the
seroprevalence of T. gondii between meat eating marsupials and herbivorous marsupials
was compared.
61
3.2.Materials and methods
3.2.1. Western grey and eastern grey kangaroos
Two hundred and nineteen western grey kangaroo blood samples were obtained from 7
different locations on the outskirts of the Perth metropolitan area, WA (Figure 3.1) over a 2
year period from May 2005 to May 2007. Wild kangaroos were culled during Department
of Environment and Conservation (DEC) population control programmes in areas such as
parks, reserves, golf courses and farms. During culling programmes an ID was allocated to
each animal and the sex of the kangaroo noted. Blood was collected by needle aspiration of
the heart within 4 hours of death of the kangaroo. Sera was separated by centrifugation and
stored at -20oC.
Eastern grey kangaroo serum samples were provided by collaborators at the University of
Queensland. One hundred and twelve blood samples were collected over a 12 month period
from 2004 to 2005 from kangaroos located in Roma, QLD (Figure 3.1). An additional 65
serum samples from eastern grey kangaroos were provided by collaborators at Macquarie
University, NSW. Blood samples were collected in May 2006 from Sydney, NSW (Figure
3.1). Western grey kangaroo and eastern grey kangaroo serum samples were tested for anti-
T. gondii IgG using the ELISA protocol outlined in section 2.2.4.
3.2.2. Woylies
In March 2006, 153 blood samples were obtained from a population of wild woylies in the
Upper Warren region, WA (Figure 3.1) via bleeding from the lateral tail vein, as part of
DEC trapping and sampling programs for the Woylie Conservation Research Project
62
(WCRP). The Upper Warren is a region of land bordering the town of Manjimup, WA.
Woylie blood samples were also collected from Dryandra Nature reserve, WA (n=12),
Batalling Forest, WA (n=17), Tutanning Nature Reserve, WA (n=8), Venus Bay Island, SA
(n=14) and St Peter’s Island, SA (n=72) (Figure 3.1). All blood samples collected were
separated via centrifugation and the serum removed was stored at -20°C. Serum samples
were tested using the commercially available MAT, as described in section 2.2.2.
3.2.3. Marsupials and native rodents in island populations
Forty four blood samples were obtained from free ranging marsupials and rodents located at
Faure Island Sanctuary, WA (Figure 3.1) as part of ongoing sampling programs being
conducted by the DEC. Blood samples were obtained in April 2007, from 28 burrowing
bettongs (Bettongia lesueur), 9 shark bay mice (Pseudomys fieldi), 5 banded hare wallabies
(Lagostrophus fasciatus) and 2 western barred bandicoots (Perameles bougainville). A
further 48 blood samples were obtained from marsupials and rodents located at Barrow
Island Nature Reserve, WA (Figure 3.1) as part of DEC sampling programs. Blood samples
were taken in September 2007, from 14 burrowing bettongs, 11 golden bandicoots (Isoodon
auratus), 8 western chestnut mice (Pseudomys nanus), 6 brush tailed possums (Trichosurus
vulpecula), 5 planigales (Planigale maculata), 3 spectacled hare wallabies (Lagorchestes
conspicillatus) and 1 water rat (Hydromys chrysogaster). Sera was separated via
centrifugation and stored at -20°C. Serum samples were tested using the commercially
available MAT, as described in section 2.2.2.
63
3.2.4. Chuditch
Twenty three blood samples were obtained from free ranging Chuditch located in Julimar
State Forest, WA (Figure 3.1) as part of DEC sampling programs. Blood samples were
obtained in June 2007 and sera was separated via centrifugation and stored at -20°C. Serum
samples were tested using the commercially available MAT, as described in section 2.2.2.
3.2.5. Statistics
The Fisher’s exact test , Chi squared test (Martin et al., 1987) or odds ratios (Dohoo et al.,
2003) were utilized to compare the seroprevalence results. The Fisher’s exact test was used
when at least one value was less than 5 and a two tailed p value was used. The Chi squared
test was used when all values were above 5 and the Pearson p value was utilised. A p value
of less than 0.05 was considered statistically significant. Odds ratios (OR) were used when
all values were above 1 and calculated with 95% confidence intervals (CI). Odds ratio
results were classified as statistically significant when the upper and lower 95% confidence
intervals did not include 1 (Dohoo et al., 2003).
The seroprevalence results of male and female western grey kangaroos were compared. In
addition, seroprevalence data from kangaroos located in Perth, Roma and Sydney were
compared. The seroprevalence of T. gondii in chuditch (carnivore) was compared to that in
western grey kangaroos (herbivore) in Perth. A retrospective case control study was
undertaken to compare seroprevalence results of marsupials located in areas where cats are
free to roam to marsupials located in areas where cats are not present. This was in order to
determine if marsupials located in areas were cats are free to roam are more likely to be
seropositive for T. gondii. Populations of marsupials located in areas where cats are not
64
present were in Faure Island, Barrow Island, St Peter Island and Venus Bay Island.
Marsupials in this study which were located in areas where cats are free to roam are western
grey kangaroos, eastern grey kangaroos and woylies in the Upper Warren, Dryandra Nature
reserve, Batalling Forest and Tutanning Nature Reserve (Morris, personal communication,
June 25, 2008). Non-marsupials were not included in the case control study.
3.3.Results
Of the 219 western grey kangaroos sampled within the Perth metropolitan area, 15.5%
(95%CI: 10.7-20.3) were seropositive for T. gondii using the ELISA. Male kangaroos had
an overall seroprevalence of 10.9% whereas females had a seroprevalence of 21.5% (Table
3.1). This difference was statistically significant (p = 0.038; OR = 0.45, CI: 0.21, 0.97).
From the 112 eastern grey kangaroos that were sampled near Roma, QLD none were
positive for anti-T. gondii IgG using the ELISA. Out of 65 eastern grey kangaroos sampled
from Sydney, NSW two were positive for anti-T. gondii IgG (Table 3.2). Thus there was a
T. gondii seroprevalence of 3.07% (95%CI: 0.0-7.3) in eastern grey kangaroos in Sydney.
The difference in seroprevalence between western grey kangaroos in Perth and eastern grey
kangaroos in Roma was statistically significant (p< 0.0001). In addition, the difference in
seroprevalence between western grey kangaroos in Perth and eastern grey kangaroos in
Sydney was statistically significant (p< 0.005, OR=5.79, CI: 1.35, 24.79). The difference in
seroprevalence between eastern grey kangaroos in Roma and those in Sydney was not
statistically significant (p>0.05).
Anti-T. gondii antibodies were detected in nine (5.8%) woylies sampled in the Upper
Warren in March 2006 (Table 3.3). All other woylie serum samples tested, except one,
65
were classified as T. gondii seronegative and had MAT titres of <1:40. One woylie serum
sample out of 73 tested from St Peter Island was found to be positive for anti-T. gondii IgG
using the MAT (Table 3.3).
All 44 serum samples from Faure Island (Table 3.4) and 48 serum samples from Barrow
Island (Table 3.5) were negative for anti-T. gondii IgG using the MAT. Out of 23 chuditch
tested from Julimar State Forest, 3 were seropositive for T. gondii. The seroprevalence of T.
gondii in chuditch was therefore 13.0% (95%CI: 0.0-26.8). The difference in seroprevalence
between chuditch in Julimar and western grey kangaroos in the Perth Metropolitan area was
not statistically significant (p>0.05).
Based on retrospective case control study which compared the seroprevalence of T. gondii
in marsupials located in areas where cats may roam (Table 3.6) to the seroprevalence of T.
gondii in marsupials located in areas without cats (Table 3.7), it was found that marsupials
located in areas where cats may roam are 14.20 (95%CI: 1.94-103.66) times more likely to
be T. gondii seropositive, compared to marsupials located in areas without cats (Table 3.8).
This result is statistically significant
3.4.Discussion
The seroprevalence of T. gondii in western grey kangaroos in this study was found to be
15.5% (95%CI: 10.7-20.3). This is similar to the 17.7% seroprevalence in wild Tasmanian
pademelons (Johnson et al., 1988), and the 15.5% seroprevalence in free ranging bridled
nailtail wallabies (Turni and Smales, 2001). The prevalence of T. gondii antibodies in
Bennett’s wallabies was lower at 3.3% and was also lower in smaller sized marsupial
66
species such as common brushtail possum and eastern barred bandicoots which were 6.3%
(Eymann et al., 2006) and 6.7% (Obendorf et al., 1996) respectively. The moderate T.
gondii seroprevalence of 15.5% in wild western grey kangaroos confirms that kangaroos can
survive with T. gondii infection in the wild. Thirty four out of 219 kangaroos in the Perth
Metropolitan area had evidence of exposure to T. gondii. There are a number of possible
sources of T. gondii infection for these kangaroos. Felids are the only definitive host of T.
gondii and there are no native Australian felids, leaving domestic and feral cats as the only
source of T. gondii oocysts. Another possible source of T. gondii infection is vertical
transmission. Evidence for vertical transmission in marsupials to date is anecdotal
(Boorman et al., 1977; Dubey et al., 1988) however it is well established in a number of
species including sheep, mice, rats, cats and humans (Duncanson et al., 2001; Johnson,
1997; Marshall et al., 2004). Kangaroos are herbivorous, therefore T. gondii infected animal
tissue is an unlikely source of infection in the western grey kangaroos tested.
In this study of western grey kangaroos, the T. gondii seroprevalence in males was
significantly less than in female kangaroos (p=0.038). Other studies have also identified a
significantly higher T. gondii seroprevalence in female sheep and goats compared to their
male counterparts (Teshale et al., 2007; van der Puije et al., 2000). It is possible that
differences in behaviour between male and female western grey kangaroos accounts for
their different levels of exposure to T. gondii oocysts. For example female kangaroos are
able to crop short grass better than males (Newsome, 1980). Males may then be forced onto
other food and have been seen on occasions camped apart in groups (Dawson, 1995).
Females which graze close to the ground may thus be more likely to be exposed to T. gondii
oocysts in soil. Recrudescence of T. gondii infection during pregnancy and a subsequent rise
of antibody titres is also a possible reason why female kangaroos had a significantly higher
67
T. gondii seroprevalence than males (Wouda et al., 1999). A rise in anti-Neospora caninum
antibodies is known to occur during pregnancy in cattle, and may be associated with
recrudescence of N. caninum infection and vertical transmission (Conrad et al., 1993;
Haddad et al., 2005; Pare et al., 1997; Wouda et al., 1999). Evidence for vertical
transmission of T. gondii in marsupials to date is anecdotal, and further studies must be
undertaken to determine if recrudescence of T. gondii infection during pregnancy (or
lactation) and subsequent vertical transmission during chronic infection occurs in
marsupials.
The difference in seroprevalence between western grey kangaroos in Perth and eastern grey
kangaroos in Roma was statistically significant. Similarly, the difference in seroprevalence
between western grey kangaroos in Perth and eastern grey kangaroos in Sydney was
statistically significant. However, the difference in seroprevalence between eastern grey
kangaroos near Roma and those in Sydney was not statistically significant. Climatic
conditions have been proposed to play a role in the prevalence of T. gondii in marsupials
and it has been suggested that a cold moist climate is associated with a high prevalence of T.
gondii infection (Attwood et al., 1975). In addition, oocysts remain viable for longer periods
of time in a cool and moist environment (Yilmaz and Hopkins, 1972). Roma, QLD and its
surrounds do have a higher average temperature and lower average rainfall than Perth and
Sydney (BOM, 2008). This difference in temperature and rainfall between Roma and Perth
may explain why kangaroos in Perth have a significantly higher T. gondii seroprevalence
than kangaroos in Roma. However the significantly higher seroprevalence of T. gondii in
Perth compared to Sydney cannot be explained by climate, as Sydney has lower average
temperatures and higher average rainfall than Perth (BOM, 2008). Therefore factors other
68
than oocyst survival may have caused the significant difference in seroprevalence between
kangaroos in Sydney and Perth.
A seroprevalence of 13.0% (95%CI: 0.0-26.8) was found in meat eating chuditch located in
Julimar State Forest. This is lower than the 20.3% seroprevalence found in Chuditch
sampled in 1993 (Haigh, 1994). The 69 chuditch tested in 1993 were sampled from Julimar
State Forest and Batalling Forest, WA. The 13.0% seroprevalence of T. gondii in chuditch
sampled in Julimar State Forest was also lower than the 15.5% seroprevalence of T. gondii
in western grey kangaroos in the Perth metropolitan area; however this difference in
seroprevalence was not statistically significant. Western grey kangaroos in the Perth
metropolitan area were the most closely located group of sampled herbivorous marsupials to
the chuditch. Julimar State Forest is approximately 65km from the centre of Perth and cats
are free to roam in both locations (Morris, personal communication, June 25, 2008). The
difference in seroprevalence between the chuditch and western grey kangaroos was not
statistically significant, therefore there was no significant difference in T. gondii
seroprevalence between meat eating marsupials and non-meat eating marsupials in this
study.
A low seroprevalence of T. gondii was found in animals located in felid-free islands. Out of
44 animals sampled from Faure Island and 48 animals sampled from Barrow Island, none
were seropositive for T. gondii. In addition, no T. gondii seropositive animals were
identified in Venus Bay Island. One serum sample from a woylie in St Peter Island was
positive for anti-T. gondii IgG out of 73 woylies sampled. It is unknown if this serum
sample gave a false positive result on the MAT or if the animal sampled was actually
infected with T. gondii. The very low seroprevalence of T. gondii in marsupials on St Peter
69
Island may be due to contamination of the environment with oocysts brought by people
from the mainland. The very low prevalence of T. gondii seropositive animals in the felid
free islands tested suggests cats play an important role in the transmission of T. gondii.
A retrospective case control study was used to determine if being located in areas where cats
are free to roam is a risk factor for T. gondii seropositivity in marsupials. Based on a number
of sample sets from a range of marsupial species it was calculated that marsupials located in
an area where cats are free to roam are 14.20 times more likely to be T. gondii seropositive,
compared to marsupials located in areas without cats. This result is consistent with other
studies which found epidemiological evidence of cats playing a role in the transmission of
T. gondii (Dubey et al., 1997a; Frenkel and Ruiz, 1981; Munday, 1972; Wallace et al.,
1972). Species difference is a confounding variable which may affect the significance of the
results. For example, several of the T. gondii seropositive marsupials in this study were
kangaroos, and no kangaroos were sampled in areas free of felids. Some marsupial species
may be more susceptible to toxoplasmosis than others and die of acute toxoplasmosis before
IgG can be detected (Johnson et al., 1988). Therefore marsupial species more sensitive to
toxoplasmosis would be expected to have a lower prevalence of anti-T. gondii IgG than
marsupial species less sensitive to toxoplasmosis. It is unknown how susceptible to
toxoplasmosis each marsupial species in the case control study is and to therefore obtain an
accurate comparison on if certain species sampled are more susceptible that others.
Infection with T. gondii in marsupials has the potential to progress to fulminant disease,
alters the way marsupials should be managed in captivity and is a public health issue as
kangaroo meat is now consumed by humans and domestic felids (Robson et al., 1995). In
this study we found both western grey kangaroos in Perth and eastern grey kangaroos in
70
Sydney and rural Queensland to be seropositive for T. gondii infection. Both western grey
kangaroos and eastern grey kangaroos are harvested for meat in Australia (Holds et al.,
2008). Kangaroo meat is sold for human and pet consumption in Australia and is also
exported to Asia, Europe and North America for human consumption (Holds et al., 2008). T.
gondii-infected kangaroos that are used for meat are a potential source of infection for
humans and also domestic cats, which may subsequently shed oocysts and perpetuate the
life cycle. Additional data comparing T. gondii seroprevalence levels between male and
female western grey kangaroos illustrates female western grey kangaroos have a
significantly higher seroprevalence rate than males, which is possibly associated with their
different levels of exposure to oocysts. Females, which feed closer to the ground than males,
are more likely to be exposed to oocysts in soil. Alternatively, recrudescence of T. gondii
infection during pregnancy/lactation may have contributed to the significantly higher T.
gondii seroprevalence in female western grey kangaroos compared to males. A low
seroprevalence of T. gondii was found in felid free island populations of marsupials. A
subsequent case control study which compares the seroprevalence of T. gondii in marsupials
located in felid-free islands to those located in areas where felids may roam found that
marsupials are more likely to be seropositive if located in areas where felids may roam.
71
Table 3.1
Prevalence of anti-T. gondii IgG in western grey kangaroos in Perth, WA as determined by
an ELISA
Sex Positive Number tested Prevalence
Male 11 101 10.89%a
Female 23 107 21.50%b
Unknown 0 11 0.00%
TOTAL 34 219 15.53%
Note: a is significantly less than b (p= 0.038)
Table 3.2
Prevalence of anti-T. gondii IgG in eastern grey kangaroos as determined by an ELISA
Location Positive Number tested Prevalence
Roma, QLD 0 112 0%
Sydney, NSW 2 65 3.07%
72
Table 3.3
Prevalence of anti-T. gondii IgG in woylies in Australia as determined by the MAT
Location Positive Number tested Prevalence
Upper Warren, WA 9 153 5.88%
Dryandra, WA 0 12 0%
Tutanning, WA 0 8 0%
Batalling, WA 0 17 0%
St Peter Island, SA 1 73 1.37%
Venus Bay Island, SA 0 14 0%
TOTAL 10 277 3.61%
Table 3.4
Prevalence of anti-T. gondii IgG in animals in Faure Island as determined by the MAT
Species Positive Number tested Prevalence
Burrowing bettong 0 28 0%
Shark Bay mouse 0 9 0%
Banded hare wallaby 0 5 0%
Western barred bandicoot 0 2 0%
TOTAL 0 44 0%
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Table 3.5
Prevalence of anti-T. gondii IgG in animals in Barrow Island as determined by the MAT
Species Positive Number tested Prevalence
Burrowing bettong 0 14 0%
Brush tail possum 0 6 0%
Golden bandicoot 0 11 0%
Planigale 0 5 0%
Western chestnut mouse 0 8 0%
Spectacled hare wallaby 0 3 0%
Water rat 0 1 0%
TOTAL 0 48 0%
Table 3.6
Combined data of anti-T. gondii IgG in marsupials located in areas where cats may roam
Location Species Positive Negative
Perth, WA Western grey kangaroo 34 185
Roma, QLD Eastern grey kangaroo 0 112
Sydney, NSW Eastern grey kangaroo 2 63
Upper Warren, WA Woylie 9 144
Dryandra, WA Woylie 0 12
Tutanning, WA Woylie 0 8
Batalling, WA Woylie 0 17
TOTAL 45 541
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Table 3.7
Combined data of anti-T. gondii IgG in marsupials located in areas without cats
Location Species Positive Negative
Faure Island, WA Burrowing bettong 0 28
Faure Island, WA Banded hare wallaby 0 5
Faure Island, WA Western barred bandicoot 0 2
Barrow Island, WA Burrowing bettong 0 14
Barrow Island, WA Brush tail possum 0 6
Barrow Island, WA Golden bandicoot 0 11
Barrow Island, WA Planigale 0 5
Barrow Island, WA Spectacled hare wallaby 0 3
St Peter Island, SA Woylie 1 72
Venus Bay Island, SA Woylie 0 14
TOTAL 1 160
75
Table 3.8
The effect of being located in an area where cats may roam on T. gondii seropositivity in
Australian marsupials
Factor Positive Negative
Percent
positive OR
Lower
95% CI
Upper
95% CI
Located in an area
where cats may roam 48 541 8.15% 14.20 1.94 103.66
Located in an area
without cats 1 160 0.62% 1.00
OR- Odds ratioCI- Confidence interval
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Figure 3.1Locations of marsupials sampledfor anti-T. gondii IgG in Australia(map obtained fromhttp://www.ga.gov.au/build/img/outline.gif)
Key to sampling abbreviations:P = Perth, WAM = Manjimup (Upper Warren), WAB = Batalling forest, WAD = Dryandra nature reserve, WAT = Tutanning nature reserve, WAJ = Julimar State Forest, WABI = Barrow Island, WAFI = Faure Island, WASPI = St Peter Island, SAVBI = Venus bay Island, SAR = Roma, QLDS = Sydney, NSW
77
4. Vertical transmission of T. gondii in Australian marsupials
4.1.Introduction
Vertical (transplacental or transmammary) transmission of T. gondii is traditionally thought
to occur infrequently and almost always in acutely infected pregnant females (Dubey and
Beattie, 1988). The influence of vertical transmission on the maintenance of T. gondii in
natural populations has been a matter of debate in recent years (Johnson, 1997). Early
studies in mice and guinea pigs found that congenital infection with T. gondii can occur
while the dam is chronically infected with T. gondii (Remington et al., 1961). This method
of vertical transmission is described as endogenous transplacental infection (TPI) (Trees and
Williams, 2005). Endogenous TPI is one of the major forms of transmission used by a
parasite very closely related to T. gondii, Neospora caninum (Dubey and Lindsay, 1996).
Recent studies verified the high frequency of congenital transmission of T. gondii in
chronically infected mice, and it was proposed that endogenous TPI can maintain T. gondii
infection in wild mice populations (Marshall et al., 2004; Owen and Trees, 1998). In
addition, a high frequency of congenital T. gondii infection was observed in naturally
infected sheep in which the resultant lambs were healthy (Duncanson et al., 2001). Recent
data also suggests T. gondii can be transmitted via successive vertical transmission within
families of sheep (Morley et al., 2005). However, studies in rats observed that congenital
toxoplasmosis, although common in acutely infected rats, is extremely uncommon in
chronically infected rats (Dubey et al., 1997b; Zenner et al., 1993). Further studies need to
be undertaken to determine the incidence of vertical transmission in other chronically
infected animals. If vertical transmission of T. gondii does occur in several species of
chronically infected animals and the resultant offspring are healthy, this would suggest that
78
vertical transmission is a more common source of T. gondii infection that previously
thought.
Evidence for vertical transmission in marsupials to date is anecdotal (Boorman et al., 1977;
Dubey et al., 1988), and the incidence of vertical transmission in marsupials is unknown.
However, considering the potential impact of toxoplasmosis in marsupials and the current
efforts associated with wildlife conservation, it is important to examine the causes of
infection of Australian marsupials with T. gondii. In some areas, the seroprevalence of T.
gondii in wild herbivorous marsupial species is as high as 17.7%, with up to 22.7% of
juveniles being positive (Johnson et al., 1988). While it is plausible that the relatively high
prevalence of T. gondii found in some populations of marsupials is due solely to
environmental contamination with oocysts from cats, it is also possible that vertical
transmission plays a role in the maintenance of T. gondii infection in marsupials.
Information on the frequency of vertical transmission in marsupials will benefit captive
breeding programmes of Australian marsupials by ensuring only T. gondii-free animals are
bred, thereby improving animal health and assisting animal conservation.
In order to better understand T. gondii transmission in marsupials, western grey kangaroos
(Macropus fuliginosus), agile wallabies (Macropus agilis) and woylies (Bettongia
penicillata) were tested for evidence of vertical transmission of T. gondii. All pouch young
in this study were tested before or close to the time of first pouch exit. Marsupial young are
born at a very immature state (less than 1gram neonatal weight) at which time they enter the
pouch. Young first exit the pouch after a long period of permanent residence (Tyndale-
Biscoe and Renfree, 1987). While within the pouch, young are protected from the external
environment and are extremely unlikely to be exposed to T. gondii oocysts.
79
Comparative immunoblots were utilised to compare dam and pouch young sera, to
differentiate maternal antibodies from actual infection in offspring. Comparative
immunoblots have been utilised in humans (Chumpitazi et al., 1995; Gavinet et al., 1997;
Gross et al., 2000; Pinon et al., 2001; Remington et al., 1985) and cats (Cannizzo et al.,
1996) to detect neonatal T. gondii infection. Neonatal T. gondii infection was diagnosed
when an IgG reactive band(s) was present in the neonate immunoblot that was absent in the
corresponding dam immunoblot (Gross et al., 2000). There are no published reports that
mention the use of comparative immunoblots to detect T. gondii infection in marsupial
young.
The MAT (modified agglutination test) and DAT (direct agglutination test) were used to test
for anti-T. gondii IgM in sera obtained from marsupial dams. Experimental studies in
eastern grey kangaroos demonstrate that a difference in titre between MAT and DAT is
indicative of an IgM response and acute T. gondii infection (Johnson et al., 1989).
Subsequent studies in range of marsupial species have successfully used the MAT and DAT
to diagnose acute T. gondii infection (Bettiol et al., 2000a; Hartley, 2006; Lynch et al.,
1993a; Skerratt et al., 1997). When tissue samples were available, immunohistochemistry
and PCR were used to detect T. gondii organisms in the tissue of dams and their
corresponding pouch young.
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4.2.Materials and methods
4.2.1. Sample collection
Western grey kangaroo sera and tissues were collected from kangaroo dams and their pouch
young, as described for western grey kangaroo group B in section 2.2.1. Briefly, from each
dam, samples of brain, tongue and sera were obtained and stored at -20°C. In addition each
pouch young was weighed and measured to estimate its age (Poole et al., 1982). Sera was
obtained from pouch young and stored at -20°C. Samples of brain, heart, skeletal muscle,
liver, lung, small intestine, kidney and spleen from each pouch young were obtained.
Sections of each tissue were placed in 10% buffered formalin for histology and the
remaining tissue was stored at -20°C for DNA extraction.
Agile wallaby sera were obtained from a group of captive wallabies which had an outbreak
of suspected toxoplasmosis in which a number of animals died. All agile wallabies were
housed at Rockhampton Zoo, QLD. Blood was collected from all wallabies via
venipuncture of the lateral tail vein. Blood was collected from the offspring of all females as
close as possible to the time of first pouch exit, which is approximately 172 to 211 days
after birth in agile wallabies (Merchant, 1976). Blood samples were collected periodically
over a two year period from August 2005 to July 2007 (Table 4.1). All blood samples
collected were separated via centrifugation and the sera was stored at -20°C. Twelve adult
agile wallabies consisting of 6 breeding females and 6 entire males were present in the
group at the beginning of the two year period. No tissue samples were taken from any of the
agile wallabies.
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A mature female woylie with young in pouch was submitted to Murdoch University
Veterinary Hospital by the Department of Environment and Conservation (DEC) for
necropsy with a history of neurological signs. Tissue samples were placed in 10% buffered
formalin and consisted of brain, heart, skeletal muscle, lung, liver, spleen and mammary
gland. In addition, brain, heart and mammary gland tissue samples were set aside in 70%
ethanol for DNA extraction. The furless pouch young present in the pouch of the necropsied
woylie had samples of brain, heart, skeletal muscle, lung and liver removed and placed in
70% ethanol for DNA extraction. The lack of fur in the woylie pouch young sampled
indicated it was too immature to have ever left the pouch.
4.2.2. Serology
The 62 western grey kangaroo dams and the corresponding 62 pouch young were all
screened for T. gondii antibodies using an ELISA to detect IgG antibodies to T. gondii in
macropod marsupials, as described in section 2.2.4. Sera from agile wallabies and their
pouch young were screened for T. gondii IgG using the commercially available MAT. The
protocol used for testing sera with the MAT is outlined in section 2.2.2. Sera samples of
seropositive western grey kangaroo and agile wallaby dams were sent to the Animal Health
Laboratory, Tasmania to obtain MAT and DAT titres in order to determine the presence of
T. gondii IgM. No serum samples were taken from the adult woylie tested or its pouch
young.
82
4.2.3. Immunoblotting
Sera from seropositive dams and their corresponding pouch young were then
immunoblotted in order to compare banding patterns of dam-young pairs. Serum samples
taken from agile wallaby dams Ag3 and Ag4 on the 11/10/2005 were tested with serum
samples from their corresponding pouch young taken on the 16/2/2006. A serum sample
from agile wallaby dam Ag7 taken on the 16/2/2006 was tested with a serum sample from
its corresponding pouch young taken on the same date. Antigen for the immunoblot was
obtained from RH strain T. gondii tachyzoites grown in Vero cell culture, as described in
section 2.2.3. T. gondii antigen was separated using sodium dodecyl sulfate polyacrylamide
gel electrophoresis (SDS-PAGE). An antigen suspension consisting of 100ug protein was
run on the SDS-PAGE along with molecular mass standards for 1 hour at 200V. Separated
proteins within the gel were then blotted onto PVDF transfer membrane (BioTrace PVDF
Membrane, PALL Gelman Laboratory, Ann Arbor, USA) using a semi-dry blotting machine
which was run at 15V for 30 minutes.
Membranes were incubated in 5% bovine skim milk in PBS for 30 minutes at 37°C on a
rocking platform. A washing cycle followed which was made up of rinsing in PBS for three
periods of 3 minutes. The membrane was then cut into strips 5mm in width and each strip
placed into individual wells. In each round of immunoblotting one T. gondii seropositive
and one T. gondii seronegative control of western grey kangaroo sera was included in
addition to a PBS control. Strips were incubated for 2 hours in sample sera which was
diluted 1:100 in PBS. Strips were then washed and incubated with commercially available
donkey anti-kangaroo IgG (Kangaroo IgG (h&I) antiserum, Bethyl Laboratories Inc,
Montgomery, USA) diluted 1:500 in PBS for 2 hours. After a washing cycle, an incubation
83
of HRP (horseradish peroxidase) conjugated rabbit anti-donkey IgG (Donkey Anti-Rabbit:
HRP, Affinity BioreagentsTM, Golden, USA) ensued for 1 hour. The strips underwent a final
washing cycle and were visualised using TMB stabilized substrate solution for HRP (TMB
Stabilised Substrate for Horseradish Peroxidase, Promega, Madison, USA), which was
incubated with the strips for 20 minutes at room temperature on a rocking platform. The
reaction was terminated by three 5 minute rinses with distilled water and the bands
examined.
4.2.4. DNA extraction and PCR
The brain and tongue of seropositive and seronegative western grey kangaroo dams and a
range of tissues of their offspring underwent DNA extraction (Table 4.2). DNA was also
extracted from the brain, heart and mammary gland tissue of a woylie dam and the brain,
heart, skeletal muscle, lung and liver of its pouch young. A number of methods of DNA
extraction were used for each sample, the protocols of which are described in section 2.2.6.
Water, tissues of seronegative dams and tissues of seronegative pouch young were used as
DNA extraction negative controls (Table 4.2).
Samples of DNA extracted from tissue samples underwent nested PCR amplification of the
ITS1 sequence (Nandra and Grigg, manuscript in preparation) in addition to the T. gondii
B1 gene, which was amplified using two different primer sets (Bretagne et al., 1993; Grigg
and Boothroyd, 2001). The PCR protocols undertaken are described in section 2.2.7. All
PCR products were sequenced, as described in section 2.2.7.
84
4.2.5. Histology and immunohistochemistry
Tissue fixed in 10% buffered formalin consisted of the heart, skeletal muscle, liver,
lung, small intestine, kidney and spleen of the 10 western grey kangaroo pouch young
which had seropositive dams. The brain, heart, skeletal muscle, lung, liver, spleen and
mammary tissue of a woylie dam were also fixed. Formalin fixed tissue samples were
trimmed and processed before being embedded in paraffin wax, sectioned and stained
with haematoxylin and eosin. Paraffin embedded tissues were also sectioned and
immunohistochemically stained with rabbit polyclonal antibodies to T. gondii, as
previously described by Lindsay and Dubey (1989). Brain tissue was not included in
histological analysis of western grey kangaroo pouch young due to marked autolysis,
associated with post-mortem changes. (Lindsay and Dubey, 1989)
4.3.Results
4.3.1. Serology
Of 62 western grey kangaroo dams which were screened for anti-T. gondii IgG using an
ELISA, 10 dams were seropositive for T. gondii and 7 of these dams had corresponding
seropositive pouch young (Table 4.2). The remaining 52 seronegative dams all had
corresponding seronegative pouch young. All 10 ELISA seropositive dams were also MAT
and DAT positive. MAT and DAT titres were identical in all western grey kangaroo dams,
which indicated a lack of IgM in all dam serum samples (Table 4.3).
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The results of agile wallaby serological screening are outlined in Table 4.1. Of the 12 initial
agile wallabies that were tested for anti-T. gondii antibodies using the MAT, 7 were
seropositive. Of the initial seropositive wallabies, four were female and three were male.
Three seropositive and two seronegative females bore pouch young. All pouch young from
seropositive dams were initially seropositive, however two out of three seroconverted, one
by approximately 333 days of age and the other by approximately 19 months of age. Only
one (Ag3PY) of the three offspring remained seropositive after weaning at 328 days of age.
Both pouch young from seronegative dams were seronegative. Of the adult agile wallabies,
three out of twelve wallabies were found to have seroconverted from positive to negative
over the two year monitoring period. All three seropositive agile wallaby dams had identical
MAT and DAT titres (Table 4.4), which indicated a lack of T. gondii specific IgM in these
animals at the time of sampling.
4.3.2. Immunoblotting
Bands against T. gondii antigens were present in immunoblots of all seropositive agile
wallaby dams and their corresponding pouch young (Figure 4.1). In two out of three initially
seropositive pouch young, all bands present were also present in the immunoblot of the
corresponding dam. One pouch young (Ag3PY) immunoblot had bands present against 18,
23 and 39kd antigens which were not present in its corresponding dam.
Bands against T. gondii antigens were also present in immunoblots of all 10 seropositive
western grey kangaroo dams and were also present in immunoblots of their seropositive
offspring (Figure 4.2). Bands were also present in 2 offspring (PYF19 and PYR19) which
were seronegative on the ELISA. All bands present in each pouch young immunoblot were
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present in the immunoblot of its corresponding dam. Consequently all pouch young tested
had antigen-antibody binding patterns consistent with passive transfer of antibody. None of
the T. gondii specific bands present in the western grey kangaroo pouch young immunoblots
could be attributed to vertical transmission of T. gondii.
4.3.3. PCR
T. gondii specific DNA was detected in all 9 seropositive western grey kangaroo dams
tested using PCR. One seropositive kangaroo dam was not tested for T. gondii DNA due to
the absence of tissue samples. No T. gondii specific DNA was detected in the DNA
extraction negative controls, which comprised tissues from three seronegative dams and 6
pouch young from seronegative dams (Table 4.2). All samples were tested with nested
primers for the ITS1 (Nandra and Grigg, manuscript in preparation) sequence and two sets
of primers for the B1 gene, one nested (Grigg and Boothroyd, 2001) and one non-nested
(Bretagne et al., 1993). DNA for T. gondii was detected in the heart tissue of two pouch
young from seropositive dams (Figure 4.3 and 4.4). No T. gondii specific DNA was
detected in tissues from the remaining 8 pouch young with seropositive dams.
PCR using primers for B1 and ITS1 detected T. gondii DNA in the mammary gland of the
one woylie tested and in the brain of its corresponding pouch young (Table 4.5). All other
woylie tissue tested was negative for T. gondii DNA. All PCR bands sequenced had
sequences that were specific for T. gondii.
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4.3.4. Histology and immunohistochemistry
Generalised congestion and oedema of the lung, likely associated with post mortem
changes, was observed in all western grey kangaroo pouch young. No other significant
lesions were observed in histological sections of 10 pouch young tested from seropositive
dams. Upon immunohistochemistry, no T. gondii tachyzoites or cysts were found despite
staining of T. gondii in positive control tissue used.
T. gondii was not detected upon histology or immunohistochemistry in the adult woylie.
Histologically there was diffuse pulmonary congestion and oedema, and autolysis of the
intestines. In addition, there was a small haematoma and focus of inflammation associated
with one mammary gland.
4.4.Discussion
The presence of T. gondii DNA was identified in the heart tissue of two pouch young from
seropositive western grey kangaroo dams and in the brain of a woylie pouch young. DNA
was also identified in the mammary gland of the woylie dam suggesting that infection of the
woylie pouch young was from suckling milk from the mammary gland. One out of three
agile wallaby pouch young from seropositive dams had an immunoblot antigen-antibody
binding pattern suggestive of actual infection with T. gondii.
It is highly unlikely that the western grey kangaroo and woylie pouch young tested in this
study were exposed to T. gondii oocysts from the external environment. Marsupial young
first exit the pouch after a long period of permanent residence and while within the pouch,
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young are protected from the external environment (Tyndale-Biscoe and Renfree, 1987). All
western grey kangaroo pouch young in this study were tested before the time of first pouch
exit, which is approximately 298 days in this species (Tyndale-Biscoe and Renfree, 1987).
In addition, the woylie pouch young in this study was unfurred and therefore too young to
have ever left the pouch. All agile wallaby pouch young from seropositive dams were tested
for T. gondii as close as possible to the time of first pouch exit, which ranges from 172 to
211 days of age in agile wallabies (Merchant, 1976). Therefore the agile wallaby pouch
young had several days contact with the external environment and the possibility of the
agile wallaby pouch young being exposed to T. gondii oocysts cannot be ruled out.
Results from the study of agile wallaby comparative immunoblots suggest one (Ag3PY) out
of three pouch young from seropositive dams was infected with T. gondii. Pouch young
Ag3PY had an antigen-antibody binding pattern suggestive of congenital T. gondii infection
whereas the other two pouch young had antigen-antibody binding patterns consistent with
passive transfer of antibodies. However, serological monitoring of the dam of Ag3PY
demonstrated that the dam’s antibody titres were waning. By the 16/2/2006, when a serum
sample was available from Ag3PY, dam Ag3 was seronegative for T. gondii using the MAT
and the serum sample taken from Ag3 from that date could not be used for comparative
immunoblot. Dam Ag4 also had waning MAT titres similar to dam Ag3. As dam Ag3 had
waning titres when its young was sampled, the comparative immunoblot results were not
reliable. Results from the agile wallaby study suggest that comparative immunoblots are not
a reliable method of diagnosing T. gondii in marsupial young. It is unknown what caused
the waning in T. gondii antibodies in agile wallaby dams Ag3 and Ag4. Comparative
immunoblots result in the western grey kangaroo samples also did not correlate with PCR
results of western grey kangaroo pouch young, which supports results from the agile
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wallaby immunoblots that comparative immunoblots are not a reliable method of diagnosing
T. gondii infection in marsupial young.
T. gondii infection was confirmed via PCR in two pouch young from seropositive western
grey kangaroo dams and one pouch young from a PCR positive woylie dam. PCR positivity
is likely to have a high correlation with actual infection with T. gondii in this study as all 9
PCR positive kangaroo adults were seropositive and all 3 PCR negative adults were
seronegative. In addition, the 6 seronegative kangaroo pouch young from seronegative dams
were PCR negative. As PCR positivity had a high correlation with actual infection in this
study it is likely that the 6 seropositive, PCR negative western grey kangaroo pouch young
were not infected with T. gondii and were only seropositive due to the passive transfer of
antibodies from the dam. Immunoblots of pouch young F19PY and R19 PY demonstrated
bands that were not present in immunoblots of the seronegative control serum, which
suggests pouch young F19PY and R19PY had low anti-T. gondii IgG titres that were not
detected in the ELISA. Pouch young F19PY and R19PY were both from seropositive dams,
and immunoblot results from these pouch young suggest they too had a passive transfer of
anti-T. gondii antibodies from their dam. The passive transfer of T. gondii specific
antibodies has also been speculated to occur in a case study of great grey kangaroos (Miller
et al., 2003). This group of kangaroos had a history of acute juvenile mortality, with death
occurring shortly after the joeys left the pouch but were still being nursed. T. gondii
seropositive dams were detected in the group using the MAT. The offspring of seropositive
dams were also tested using the MAT. It was found that MAT titres in all 6 juvenile
kangaroos decreased over time. Miller et al (2003) argued that the decreasing T. gondii titre
in juvenile kangaroos was indicative of decreasing maternal antibodies and ruled out actual
T. gondii infection in the juvenile kangaroos. However, it was not possible to confirm the
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presence of T. gondii infection in any of these juvenile kangaroos as all survived making no
tissue samples available for analysis using histology or PCR. Due to the lack of tissue
samples in Miller et al (2003), it can be argued that T. gondii titres decreased in some of the
juvenile kangaroos due to natural reduction in IgG titres after initial T. gondii infection,
rather than due to passive immunity.
The detection of T. gondii DNA in only the heart muscle of the 2 western grey kangaroo
pouch young is not surprising as T. gondii has been detected previously in the heart muscle
of a black-faced kangaroo (Macropus fuliginosus melanops) pouch young (Dubey et al.,
1988) and two juvenile common wombats (Vombatus ursinus) (Hartley, 2006), and is
known to commonly infect the heart of adult macropod marsupials (Basso et al., 2007;
Canfield et al., 1990; Lynch et al., 1993a; Reddacliff et al., 1993). However, no pathology
or T. gondii organisms were observed in histological sections of pouch young tissue from
western grey kangaroos, despite immunohistological staining. This suggests neither of the
pouch young tested, both of which were from seropositive dams, had clinical toxoplasmosis.
In a study of tammar wallabies, no histological lesions consistent with toxoplasmosis were
observed in two out of nine experimentally infected wallabies (Reddacliff et al., 1993). Both
wallabies without histological lesions were asymptomatic for T. gondii infection, with the
remaining having severe clinical signs of toxoplasmosis. The observation of PCR positive,
histologically negative pouch young in this study therefore suggests they had asymptomatic
T. gondii infection. Since T. gondii asymptomatic offspring are likely to survive until
adulthood, such congenital transmission could contribute to the prevalence of T. gondii
infection in the wild population of kangaroos in this study.
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The mammary gland of a woylie dam and the brain of its unfurred pouch young were also
PCR positive. This is suggestive of milk transmission of T. gondii from the woylie dam to
the pouch young via the mammary gland. T. gondii in the milk is the likely mechanism of
vertical transmission in marsupials as opposed to transplacental infection. This is because
marsupial young are born at a very immature state and milk is the source of nourishment
which enables young to develop to a stage where they can leave the pouch. The average
weight of young at birth is a tiny 290 mg in woylies, 630 mg in agile wallabies and 828 mg
in western grey kangaroos. Therefore if pouch young are infected in-utero in this immature
state, they are not likely to survive past initial infection. Vertical transmission via the milk
was recently proposed by Johnson (1997) to have a role in the life cycle of T. gondii. Milk
transmission of T. gondii is not well documented, however tachyzoites have been isolated
from the milk of a number of species including mice, cats, cows, pigs, dogs, sheep, rats,
guinea pigs and rabbits (Johnson, 1997). Tachyzoites are orally infective to cats and mice
(Dubey, 1998) which suggests tachyzoites in milk are infective via the gastrointestinal
route. In addition, acid-resistant T. gondii bradyzoites were found in the milk of lactating
mice and able to produce consistent infection via the gastrointestinal route (Pettersen, 1984).
Several studies have suggested congenital transmission via the milk is common in cats
(Dubey, 1995; Powell et al., 2001; Powell and Lappin, 2001). Humans have also been
infected with T. gondii by drinking unpasteurised goat’s milk (Riemann et al., 1975; Sacks
et al., 1982) and transmission of T. gondii in humans through breastfeeding has been
suspected (Bonametti et al., 1997).
No T. gondii-specific IgM antibodies were detected in any of the 10 seropositive western
grey kangaroo dams or 3 seropositive agile wallaby dams. Each dam had a high DAT titre
which correlated exactly with its MAT titre and this illustrates a lack of IgM in the sera
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samples. The lack of IgM in the sera samples is suggestive of chronic T. gondii infection in
the western grey kangaroos and agile wallabies. Of 10 chronically infected western grey
kangaroos, 2 had T. gondii-positive offspring. This result adds support for the role of
vertical transmission in the maintenance of T. gondii infection in marsupial populations, as
it is not only acutely infected marsupials that have T. gondii-infected offspring.
The frequency of congenital transmission found in this study of marsupials is lower than in
studies of naturally infected mice (Marshall et al., 2004) and sheep (Duncanson et al., 2001;
Williams et al., 2005). However, one other study detected a relatively low rate (4.1%) of
congenital transmission in seropositive sheep using serology, PCR and histology (Rodger et
al., 2006). Further studies, which test larger numbers of young from T. gondii infected
animals need to be undertaken in order to obtain a significant comparison of congenital
infection rates between marsupials and other species.
Of two published cases of suspected congenital T. gondii infection in marsupials, all three
pouch young died and had severe T. gondii associated pathology (Boorman et al., 1977;
Dubey et al., 1988). This is different from the two western grey kangaroo pouch young in
this study which did not have observed T. gondii related pathology despite being infected
with T. gondii. It is highly unlikely that PCR results from the two PCR positive western
grey kangaroo pouch young were false positive as all negative controls used, including
water controls and tissue from seronegative kangaroos were PCR negative. Reasons for
differences in pathology between different cases of suspected congenital T. gondii infection
include species differences, maturity of young when infected, T. gondii strain type, and the
presence of stressors such as captivity or poor animal management.
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The varied results of studies which have tested for congenital T. gondii infection indicate
that the transfer of T. gondii from mother to offspring is dependent on several factors. One
factor which has been confirmed as having an effect on the incidence of congenital T. gondii
infection in humans is timing of infection in utero (Dunn et al., 1999). In addition, the
timing of infection in utero is inversely related to the severity of disease in congenitally
infected humans (Montoya and Liesenfeld, 2004). Additional factors in animals that have
been speculated to influence the probability of congenital infection with T. gondii include
the size of the placenta, length of gestation, immunocompetence of the foetus and maternal
immunity (Johnson, 1997). In Europe, type II strains of T. gondii tend to be found more
commonly in cases of congenital T. gondii infection, and it has been suggested that the
genotype of T. gondii affects the chance and severity of congenital infection (Darde et al.,
2007). The genotype of T. gondii infecting a western grey kangaroo pouch young is
investigated in the following chapter.
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Table 4.1
MAT results from agile wallabies and their offspring
ID Sex 9/08/2005 11/10/2005 16/02/2006 1/06/2006 10/07/2007
Ag1 F Positive Positive Positive Positive Positive
Ag2 M Negative Negative Negative Negative nd
Ag3 F Positive Positive Negative nd nd
Ag4 F Positive Positive Negative Negative Negative
Ag5 F Negative Negative Negative Negative nd
Ag6 F Negative Negative nd nd nd
Ag7 F Positive Positive Positive Positive Positive
Ag8 M Negative Negative Negative Negative nd
Ag9 M Positive Positive Positive Positive Positive
Ag10 M Negative Negative nd nd nd
Ag11 M Positive Negative nd nd nd
Ag12 M Positive Positive nd nd nd
Ag3PY F nd nd 100 days Positive 228 days Positive 333 days nd
Ag4PY U nd nd 100 days Positive 228 days Negative 333 days Negative 737 days
Ag5PY U nd nd 80 days nd 202 days Negative 313 days nd
Ag6PY F nd Negative 210 days nd nd nd
Ag7PY M nd nd 60 days Positive 188 days Positive 293 days Negative 697 daysU- Unknownnd- No sample availablePY- Pouch youngdays- number of days old
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Table 4.2
ELISA and PCR results from western grey kangaroo dams and their pouch young
PCR Dams PCR PY
Dam IDELISADam
ELISAPY
Age PY(days) Brain Tongue Brain Heart Sk musc Lung Liver Kidney Spleen Sm intest
C14 Positive Positive 145 B1, ITS1 Negative Negative B1 Negative Negative Negative nd Negative Negative
C9 Positive Positive 114 B1, ITS1 ITS1 Negative Negative Negative Negative Negative nd Negative Negative
J6 Positive Positive 124 B1, ITS1 ITS1 Negative Negative Negative nd nd nd nd Negative
J10 Positive Positive 125 nd B1, ITS1 Negative Negative Negative Negative Negative Negative Negative Negative
R7 Positive Positive 90 B1, ITS1 Negative Negative Negative Negative nd nd nd nd nd
Q1 Positive Positive 154 Negative B1, ITS1 Negative Negative Negative nd nd nd nd nd
G21 Positive Negative 58 B1 ITS1 Negative Negative Negative nd nd nd nd nd
F19 Positive Negative 132 ITS1 Negative Negative Negative Negative nd nd nd nd nd
R19 Positive Negative 142 ITS1 Negative Negative B1, ITS1 Negative nd nd nd nd nd
15B1 Positive Positive 246 nd nd Negative Negative Negative Negative Negative Negative Negative Negative
R4 Negative Negative 89 nd nd Negative Negative nd nd nd nd nd nd
F8 Negative Negative 98 Negative Negative Negative Negative nd nd nd nd nd nd
H14 Negative Negative 84 Negative Negative Negative Negative Negative nd nd nd nd nd
I14 Negative Negative 75 Negative Negative Negative Negative nd nd nd nd nd nd
Q20 Negative Negative 129 nd nd Negative Negative nd nd nd nd nd nd
15B2 Negative Negative 233 nd nd Negative Negative Negative Negative Negative Negative Negative Negative
PY- Pouch youngB1- Positive B1 PCR (Bretagne et al., 1993)B1- Positive B1 PCR (Grigg and Boothroyd, 2001)ITS1- Positive ITS1 PCR (Nandra and Grigg, manuscript in preparation)Negative- Negative on all PCRsnd- No test undertakenSk musc- Skeletal muscleSm intest- Small intestine
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Table 4.3
T. gondii DAT and MAT titres of seropositive western grey kangaroo dams
Dam ID DAT MAT
C14 256000 256000
C9 256000 256000
J6 4096 4096
J10 4096 4096
R7 64000 64000
Q1 4096 4096
G21 64000 64000
F19 4096 4096
R19 4096 4096
15B1 64000 64000
Table 4.4T. gondii DAT and MAT titres of seropositive agile wallaby dams
9/08/2005 11/10/2005
Dam ID DAT MAT DAT MAT
Ag3 64 64 256 256
Ag4 16000 16000 nd nd
Ag7 64000 64000 64000 64000nd- No serum available
Table 4.5T. gondii PCR results of a woylie dam and its pouch young
Tissue Woylie Dam Woylie PY
Brain Negative B1, B1, ITS1
Heart Negative Negative
Mammary gland B1, B1, ITS1 nd
Skeletal muscle nd Negative
Lung nd Negative
Liver nd NegativePY- Pouch youngB1- Positive B1 PCR (Bretagne et al., 1993)B1- Positive B1 PCR (Grigg and Boothroyd, 2001)ITS1- Positive ITS1 PCR (Nandra and Grigg, manuscript in preparation)Negative- Negative on all PCRsnd- No sample available
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Figure 4.1 Comparative immunoblots of seropositive agile wallaby dams and theiryoung.Arrows point to bands in pouch young immunoblot that are not present in the damimmunoblot.PY-Pouch youngN- seronegative control consisting of an MAT negative agile wallaby serum samplePBS- PBS control
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Figure 4.2 Comparative immunoblots of seropositive western grey kangaroo dams andtheir pouch young.PY-Pouch young
99
M 1 2 3 4
Figure 4.3 Non-nested B1 PCR of western grey kangaroo tissue DNALane M, 100bp DNA ladder; Lane 1, pouch young C14 heart tissue; Lane 2, adult G21 braintissue; Lane 3, T. gondii RH strain positive control; Lane 4, water negative control
Figure 4.4 Nested B1 PCR of western grey kangaroo tissue DNALane M, 100bp DNA ladder; Lane 1, pouch young R19 heart tissue; Lane 2, pouch youngR19 heart tissue 1:10 dilution; Lane 3, T. gondii RH strain positive control; Lane 4, waternegative control
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5. Molecular characterization of T. gondii isolates from
Australia
5.1.Introduction
It is widely thought that T. gondii has a low genetic diversity due to the common finding of
strains that can be grouped into three highly clonal but closely related lineages (Howe and
Sibley, 1995; Johnson, 1997; Su et al., 2003). However, it is increasingly being proposed
that the genetic diversity among T. gondii strains is greater than current estimates due to the
sampling bias that has resulted from the study of strains from humans and domestic animals
primarily originating from North America and Europe (Ajzenberg et al., 2004). T. gondii
isolates from wildlife or isolated parts of the world are commonly found to have atypical
genotypes (Darde et al., 2007). These atypical isolates fall into two general classes:
‘recombinant’ strains which have genotypes that are clearly related to the three dominant
types; and ‘novel’ strains which have many unique polymorphisms and novel alleles.
Atypical isolates were sampled mainly from geographically isolated regions in French
Guiana (Bossi et al., 1998; Carme et al., 2002; Darde et al., 1998) and Brazil (Ferreira et al.,
2006; Khan et al., 2006) or in wildlife such as deer, bear, cougar or sea otter (Darde et al.,
1998; Howe and Sibley, 1995; Lehmann et al., 2000; Miller et al., 2004).
A number of studies which have genotyped T. gondii isolates from wildlife (Dubey et al.,
2004a) and from geographically isolated locations (Dubey et al., 2002; Dubey et al., 2003a;
Dubey et al., 2003b; Dubey et al., 2003c; Dubey et al., 2003d) have not found atypical
strains. Many of these other studies have used a limited number of polymerase chain
reaction restriction fragment length polymorphism (PCR-RFLP) markers to identify isolates
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as either type I, II or III. Studies which have identified atypical strains of T. gondii (Bossi et
al., 1998; Carme et al., 2002; Darde et al., 1998; Howe and Sibley, 1995; Lehmann et al.,
2000) have often used techniques including isoenzyme analysis, microsatellite analysis and
multilocus genotyping. PCR-RFLP analysis using as many as three loci may at times
misidentify atypical strains as belonging to one of the three clonal lineages (Grigg et al.,
2001b). Therefore, it is possible that the many studies to date which have genotyped T.
gondii isolates using a small number of PCR-RFLP genetic markers have overlooked
atypical strains.
Different strains of T. gondii have been linked with different clinical outcomes in humans
and animals. Type I strains are highly virulent in mice and are also associated with severe
ocular toxoplasmosis in immunocompetent humans (Boothroyd and Grigg, 2002; Grigg et
al., 2001b; Vallochi et al., 2005). Atypical type X strains have also been associated with
severe meningoencephalitis in Californian sea otters (Enhydra lutris nereis) (Miller et al.,
2004). To date, there have been no studies published that molecularly characterized T.
gondii isolates from Australia. Considering previous data which suggests T. gondii isolates
from wildlife or geographically isolated areas are more likely to have atypical genotypes,
genotypic analysis of T. gondii isolates from Australian wildlife is of special interest. In
addition to its extreme isolation, Australia hosts a myriad of remarkable wildlife species.
Australian marsupials in particular are known for their unusual susceptibility to manifest T.
gondii related disease. Infection with T. gondii has been responsible for numerous deaths in
captive marsupials (Barrows, 2006; Boorman et al., 1977; Canfield et al., 1990; Dubey et
al., 1988; Hartley et al., 1990; Miller et al., 2000; Miller et al., 1992; Patton et al., 1986) and
has been attributed to causing declines in Australian marsupial populations in the wild
(Eymann et al., 2006; Obendorf et al., 1996). The aim of this study was to analyse the
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molecular characteristics of T. gondii isolates from Australia, particularly those from
marsupials. Sequencing of PCR products obtained using nested primers for the 35 copy B1
gene was used to identify single nucleotide polymorphisms (SNPs) in T. gondii DNA
isolates. The function of the B1 gene is not yet known (Switaj et al., 2005). PCR for the B1
gene was chosen to amplify T. gondii DNA because it is routinely used for highly sensitive
and specific detection of T. gondii DNA in clinical specimens (Grigg and Boothroyd, 2001).
The B1 gene is present in 35 copies in the T. gondii genome, making PCR targeting the B1
gene more sensitive than PCR targeting single copy loci such as GRA6 and SAG3.
5.2.Materials and methods
5.2.1. Sample collection
One hundred and twenty eight tissue samples were used to test for T. gondii DNA, which
comprised of 20 meat samples and a number of tissue samples from 40 animals (Table 5.1).
Eighteen chilled packets of mince were bought from local supermarkets in Perth, WA.
Mince samples tested for T. gondii consisted of 8 packets of kangaroo mince for human
consumption, 2 packets of kangaroo mince for pets, 5 packets of lamb mince, 1 packet of
mutton mince and 2 packets of pork mince. Packets of meat were stored at 4°C until
processed. Bradyzoites were purified from mince samples in order to maximise the
probability of finding T. gondii DNA. From each mince packet, 100 grams of meat was
sampled and purified using pepsin/HCl digestion (Gajadhar and Marquardt, 1992). Briefly,
for every 100 gram meat sample, 400ml of pepsin/HCl solution was used which consisted of
400ml of water mixed with 2.8ml of 18M HCl, 1.0 grams of pepsin (Pepsin from porcine
gastric mucosa powder, Sigma-Aldrich, Castle Hill, Australia) and 2.0 grams of NaCl. The
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meat mixture was incubated with gentle agitation for 1 hour at 37°C. Four layers of gauze
were used to remove undigested tissue and the filtrate was washed twice in warm (37°C)
PBS by centrifugation at 3000 RPM for 5 minutes. The bradyzoites present in the pellet
were purified using a Percoll gradient. The Percoll gradient was made using 5ml of
bradyzoites suspension in PBS and 10ml of Percoll (Amersham Biosciences, Uppsala,
Sweden). This solution was centrifuged at 2000 RPM for 20 minutes. Bradyzoites were
recovered from the pellet and washed twice in warm (37°C) PBS by centrifugation at 3000
RPM for 5 minutes. The resulting suspension was visualised under the microscope to check
for bradyzoites and stored at -20°C.
Animals which exhibited neurological signs or sudden death were deliberately targeted for
tissue sampling in order to maximise the probability of finding T. gondii DNA. A western
ringtail possum (Pseudocheirus occidentalis) with a history of hindlimb paralysis had tissue
samples removed to test for T. gondii DNA. Heart and skeletal muscle samples were
obtained from collaborators at Murdoch University Veterinary Hospital, WA and stored at -
20°C. Two wild woylies (Bettongia penicillata) which were found dead in the field had
brain and heart samples removed and stored at -20°C for DNA extraction. Tissue samples
from meerkats (Suricata suricatta) inhabiting Perth Zoo, Western Australia were obtained
after an outbreak of suspected toxoplasmosis in which a number of meerkats suffered
neurological signs and subsequently died. Brain samples from three meerkats exhibiting
neurological signs were obtained from Perth Zoo and stored at -20°C for DNA extraction.
Clinical records and pathology reports available from sampled animals were compiled. In
addition, samples of the feed of the meerkats were obtained. Meerkats at Perth Zoo were fed
a diet which included horse meat and whole mice. Two samples of horse meat were
obtained from Perth Zoo and stored at -20°C for DNA extraction. In addition, the brains of 4
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mice were removed under sterile conditions and stored at -20°C for DNA extraction. Sterile
water was used as DNA extraction negative controls.
A number of samples in this study were already tested for T. gondii DNA in vertical
transmission studies, outlined in section 4.2.4. and 4.3.3. These samples consisted of tissue
from western grey kangaroo and woylie dams and their offspring (Table 5.1). All PCR
products from samples tested using B1 primers (Grigg and Boothroyd, 2001) during the
vertical transmission studies were set aside for DNA sequencing.
5.2.2. DNA extraction and PCR
DNA samples were extracted using four different methods as outlined in section 2.2.6.
Samples of DNA extracted from tissue samples underwent nested PCR amplification of the
T. gondii B1 gene (Grigg and Boothroyd, 2001). Each sample of DNA was tested twice.
DNA from the RH strain was used as a PCR positive control and distilled water was used as
a negative control. Reaction mixtures and amplification conditions are outlined in section
2.2.7.
5.2.3. DNA sequencing
B1 gene PCR products that were 492bp in size were cut from 0.8% agrose gels stained with
ethidium bromide. DNA was purified from agrose gels using the UltraClean GelSpin DNA
Extraction Kit (MO BIO Laboratories Inc, Carlsbad, USA). Sequencing reactions were
performed directly on PCR products using a BigDye Terminator v3.1 Cycle Sequencing
Kit (Applied Biosystems, Scoresby, Australia) according to the manufacturer’s directions
105
and using internal primers for the B1 gene (Grigg et al., 2001b). Reactions were
electrophoresed through an ABI 3730 automatic sequencer and sequencing profiles analysed
using FinchTV version 1.4 (Geospiza, Seattle, USA). Bioedit version 7.0.0 (Ibis
Biosciences, Carlsbad, USA) was used to align the sequences for comparison. DNA samples
which were B1 PCR positive were retested using B1 PCR and the PCR products sequenced
once more to assess the reproducabilty of sequencing results.
5.3.Results
5.3.1. PCR of the B1 gene and sequencing of PCR products
Of 128 tissue samples tested, PCR products corresponding to the B1 gene of T. gondii were
found in 13 tissue samples, which comprised 11 animals and 2 meat samples (Table 5.1). B1
gene PCR products were subsequently sequenced to determine the genotype of T. gondii.
Out of 13 tissue samples sequenced at the B1 gene, 6 had a type I allele whereas 7 were had
SNPs inconsistent with strains I, II, III or X (Table 5.2). The SNPs were reproducible on
subsequent B1 gene PCR and sequencing.
Out of the 7 isolates with atypical sequences, four had a unique polymorphism at position
378, not documented in any strains to date. Isolates C9B and K2.8 had an adenosine/guanine
dinucleotide site at position 378 and isolates R7 and A13 had an adenosine SNP at position
378. Furthermore, isolate C14B had a unique adenosine/cytosine dinucleotide site at
position 533, not documented in any strains to date. Isolate C14B also had a SNP found in
type II and type III strains, which was a cytosine/thymine dinucleotide site at position 366.
However, isolate C14B did not have the other B1 gene polymorphism of type II and type III
106
strains, which was a cytosine/guanine dinucleotide site at position 504. Isolate J6B had a
unique polymorphism not documented in any strains to date, this was a thymine at position
317. In addition, isolate J6B had a cytosine present at position 504, which is a SNP shared
by the type X strain, and guanine present at position 360, which is similar to the
cytosine/guanine dinucleotide site found in the type X strain.
Five isolates had SNPs at position 504. Three of these 5 isolates with SNPs at position 504
had a cytosine/guanine dinucleotide polymorphism at position 504 (isolates C9B, K2.8 and
A13) which was identical to the polymorphism found in type II and III strains. Despite this,
none of these three isolates (C9B, K2.8 and A13) had the additional polymorphism typical
of type II and type III strains, which was a cytosine/thymine dinucleotide site at position
366. The remaining 2 isolates with an SNP at position 504, isolates J10T and J6B, had a
cytosine polymorphism, which is shared by the type X strain. However, neither of these 2
isolates shared the other B1 gene polymorphism of the type X strain, which was a
cytosine/guanine dinucleotide site at position 360. Two samples, C9B and K2.8, had
identical SNPs in the B1 gene. In total, 6 atypical genotypes were identified in Australia,
each with a different combination of SNPs.
5.3.2. Clinical history and pathology of PCR positive animals
Of the animals whose tissues were sampled, neurological signs were observed in two
meerkats, one woylie and one western ringtail possum. Of these animals, two were positive
for T. gondii DNA based on PCR of the B1 gene; meerkat A13 and woylie A1. Meerkat
A13 had an atypical strain of T. gondii that was isolated from its brain. It was a captive
meerkat and inhabited an enclosure at Perth Zoo. This meerkat exhibited an abnormal gait at
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approximately 3 years of age, which progressively worsened until the date of euthanasia 6
months later. Within one month of the date of death, the meerkat had three episodes of
seizures and became ataxic. Other clinical signs included circling behaviour and facial
twitching. No gross abnormalities were observed on pathology. Histopathology results
revealed non-suppurative encephalitis and a plasma-lymphocytic myocarditis. No T. gondii
organisms were found histologically, despite immunohistochemistry for T. gondii.
Woylie A1 had T. gondii DNA isolated from its mammary gland. It possessed a type I allele
at the B1 gene. The woylie was a wild marsupial, caught in the field using a trap. When
removed from the trap and handled the woylie started convulsing after which it suddenly
died. Significant gross lesions on necropsy were small ecchymotic haemorrhages over the
head and within the left temporal muscle. A large subdural haematoma was present in the
left cerebral hemisphere however there was no evidence of skull fractures. The lungs were
also bilaterally congested. Histologically there was diffuse pulmonary congestion and
oedema. The intestines were autolyzed. There was also a small haematoma and focus of
inflammation within one mammary gland. The final diagnosis of cause of death was an
acute subdural haematoma. No T. gondii organisms were observed in woylie A1 during T.
gondii immunohistochemistry. It is unknown if woylie A1 died of toxoplasmosis as no
histological lesions diagnostic of toxoplasmosis were found. No information on the clinical
history of the other PCR positive animals was available. Of the remaining animals which
were PCR positive, one (PYR19) was examined using histology and immunohistochemistry.
Kangaroo PYR19 had T. gondii DNA isolated from its heart muscle which had a type I
allele at the B1 gene. Histologically there was generalised congestion and oedema of the
lung with no other significant lesions. No T. gondii organisms were observed in PYR19
upon immunohistochemistry.
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5.4.Discussion
Sequencing of the B1 gene revealed atypical genotypes in 7 out of 13 samples from
Australia. These 7 isolates contained SNPs in the B1 gene that could not be matched with
known sequences from strains I, II, III and X. It is apparent the SNPs are due to true genetic
variation because PCR reactions were repeated once, with the same results. The result of
53.8% of strains from Australia being atypical is different from the results reported by
(Howe and Sibley, 1995) who studied 106 strains, predominately from Europe and North
America, and found >95% (102 out of 106) to fall into three distinct genotypes. The result
of a majority of T. gondii isolates sampled from native Australian marsupials being of an
atypical genotype suggests that T. gondii in wild marsupials in Australia does not have a
strictly clonal population structure.
Of the 10 T. gondii isolates from Australian marsupials sequenced, 6 had an atypical
genotype, with the remainder having a type I allele at the B1 gene. Five unique genotypes
were identified out of the 6 atypical isolates from Australian marsupials; two out of the 6
isolates had the same unique sequence at the B1 gene whereas the other 4 isolates each had
different combinations of SNPs at the B1 gene. The result of a majority of T. gondii isolates
from native Australian wildlife having a range of unique genotypes is similar to the findings
of Ajzenberg et al (2004) where 9 isolates sampled from French Guiana were all atypical
and each isolate had a unique genotype. Ajzenberg et al (2004) argued that the high T.
gondii genetic diversity found in French Guiana may be due high host diversity in the wild
population which drives greater T. gondii genetic diversity through enhanced sexual
propagation in felid hosts and subsequent genetic recombination. Similarly the high genetic
109
diversity of T. gondii found in native Australian marsupials may be due to the high host
diversity in the wild populations sampled which caused enhanced sexual propagation in
felid hosts and subsequent genetic recombination. There is speculation that the
predominance of clonal strains in North America and Europe is due to their production of a
small range of domestic meat-producing animals and the subsequent propagation of clonal
strains via tissue cysts (Ajzenberg et al., 2004; Darde et al., 2007; Dardé et al., 2008).
Australia’s extreme isolation from the rest of the world and strict quarantine protocols may
favour the predominance of unique recombinations of the T. gondii introduced with the first
settlers. Australia may harbour T. gondii genotypes that have evolved independently from T.
gondii present in the rest of the world. T. gondii in Australia may have evolved to create a
genotype(s) that is less virulent to Australian marsupials.
The T. gondii genome appears to be considerably conserved with a less than 2% difference
at DNA sequence level among predominant clonal genes (Ajzenberg et al., 2004). The B1
gene in particular has a low amount of polymorphisms present compared to genes such as
GRA6 and SAG2 (Fazaeli et al., 2000; Grigg and Boothroyd, 2001; Lehmann et al., 2000).
Within the B1 gene PCR product amplified, there exists only two polymorphisms to
differentiate type I strains from type II/III strains (Grigg and Boothroyd, 2001). In addition,
several publications mention the B1 gene is conserved (Jones et al., 2000; Lee et al., 2008;
Mavin et al., 2004; Pelloux et al., 1996). However, in this study we have found a relatively
high number of SNPs (1 to 3) within the B1 gene in a large percentage of isolates
sequenced. The results demonstrate that the B1 gene is polymorphic and that Australian
isolates have a different genotype to those found elsewhere to date. Although T. gondii
isolates from domestic animals such as horses and mice were analysed in this study, a more
in depth study comparing T. gondii isolates from wildlife to isolates from humans and
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domestic animals would provide important information regarding the origin and
diversification of T. gondii in Australia.
A high proportion (6 out of 13) of T. gondii isolates found in this study of Australian
animals had a type I allele at the B1 gene, whereas the rest had atypical alleles. This is
different from the situation in North America and Europe where the majority of T. gondii
isolates in animals and humans found are type II strains (Darde et al., 1992; Howe et al.,
1997; Howe and Sibley, 1995). A similar situation to Australia occurs in Brazil, where a
high proportion of isolates found have been genotyped as type I, recombinants of type I or
novel strains (Ferreira et al., 2006; Khan et al., 2006). Type I strains are known to cause
lethal infection in mice, whereas type II and III strains are relatively nonvirulent in mice
(Sibley and Boothroyd, 1992). Type I strains or strains bearing type I alleles have been
linked to cases of ocular toxoplasmosis in immunocompetent humans (Boothroyd and
Grigg, 2002; Grigg et al., 2001b; Khan et al., 2006; Vallochi et al., 2005). Speculation arose
that the high frequency of type I strains found in Brazil may be in part responsible for the
high frequency of acquired ocular toxoplasmosis in humans in Brazil (Ferreira et al., 2006;
Khan et al., 2006). Cases of ocular toxoplasmosis in Brazil are often recurrent and serious in
nature (Glasner et al., 1992; Silveira et al., 2001). The clinical significance of finding a high
proportion of strains from Australia bearing a type I allele at the B1 gene is unknown at this
stage.
Of the animals in which T. gondii DNA was found, two had a history of neurological signs;
one meerkat and one woylie. An atypical T. gondii genotype was isolated from the brain of
meerkat A13. The encephalitis and myocarditis found in meerkat A13, in combination with
the finding of T. gondii DNA in brain tissue strongly suggest that T. gondii was responsible
111
for the clinical deterioration of this animal. Several observations indicate that meerkats, like
Australian marsupials, are highly susceptible to toxoplasmosis (Juan-Sallés C et al., 1997).
A suspected outbreak of toxoplasmosis occurred in the enclosure which meerkat A13
inhabited. In the enclosure five out of 11 meerkats exhibited neurological signs and three
died. T. gondii DNA was found in two out of the three animals that died using PCR for the
ITS1 sequence (Nandra and Grigg, manuscript in preparation) and B1 gene (Grigg and
Boothroyd, 2001), however no T. gondii organisms were observed in any of the deceased
meerkats using histology. Of the two remaining live meerkats that exhibited neurological
signs, both were T. gondii seropositive using the modified agglutination test. A
toxoplasmosis outbreak has also been documented in a group of meerkats in Barcelona Zoo
in which seven out of nine meerkats died (Juan-Sallés C et al., 1997). It is unclear if the
strain of T. gondii had a direct effect on the pathology of meerkat A13. Further studies
linking T. gondii strain with T. gondii-related pathology in meerkats needs to be undertaken
to address this issue. In depth studies in sea otters found that otters infected with type X
strains tend to have moderate to severe meningoencephalitis on histopathology more
frequently than type II infected otters (Miller et al., 2004).
At this stage it is unknown if the atypical strains found in this study are more commonly
associated with certain T. gondii related disease manifestations. However the result of 7 out
of 13 T. gondii isolates from Australia bearing atypical alleles is consistent with suggestions
from Ajzenberg et al (2004) who propose that the genetic diversity of T. gondii in wildlife
and geographically isolated areas is underestimated. Further studies in Australia are
necessary to determine the prevalence of atypical strains. Additional studies linking atypical
strains with their clinical manifestation are also warranted.
112
In this study of the B1 sequence of T. gondii isolates from Australia, it was fortunate that
SNPs were found and thus atypical isolates discovered. Nevertheless, future analysis should
be undertaken on the T. gondii DNA isolates from this study using a multi-locus approach to
further characterise all isolates, both those that are atypical and those in which no SNPs
were found.
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Table 5.1
Tissue samples tested for T. gondii DNA using PCR of the B1 gene
Sample ID Animal Species AgeT. gondiiIgG
AnimalID Tissue B1 PCR Remarks
A1a Woylie Bettongia penicillata Adult nd WB2229 Brain NegativeWild woylie found withneurological signs
A1b Woylie Bettongia penicillata Adult nd WB2229Mammarygland Positive
Wild woylie found withneurological signs
A1c Woylie Bettongia penicillata Adult nd WB2229 Heart NegativeWild woylie found withneurological signs
A1Ya Woylie Bettongia penicillata PY ndPY ofWB2229 Brain Positive
Pouch young of woylieWB2229
A1Ya Woylie Bettongia penicillata PY ndPY ofWB2229 Heart Negative
Pouch young of woylieWB2229
A1Ya Woylie Bettongia penicillata PY ndPY ofWB2229
Skeletalmuscle Negative
Pouch young of woylieWB2229
A1Ya Woylie Bettongia penicillata PY ndPY ofWB2229 Lung Negative
Pouch young of woylieWB2229
A1Ya Woylie Bettongia penicillata PY ndPY ofWB2229 Liver Negative
Pouch young of woylieWB2229
A14bWestern ringtailpossum
Pseudocheirusoccidentalis Adult nd 05-971
Skeletalmuscle Negative
Wild possum found withneurological signs
A14cWestern ringtailpossum
Pseudocheirusoccidentalis Adult nd 05-971 Heart Negative
Wild possum found withneurological signs
A12 Meerkat Suricata suricatta AdultNegativeMAT 950721 Brain Negative
Captive meerkat withneurological signs, PerthZoo
A13 Meerkat Suricata suricatta Adult nd A01070 Brain Positive
Captive meerkat withneurological signs, PerthZoo
A15 Meerkat Suricata suricatta Adult nd 890004Formalinfixed brain Negative
Captive meerkat withneurological signs, PerthZoo
A6 Horse Equus caballus Unknown nd A6 Horse Negative Sample of horse meat fed
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meat to meerkats at Perth Zoo
A7 Horse Equus caballus Unknown nd A7Horsemeat Positive
Sample of horse meat fedto meerkats at Perth Zoo
A8 Mouse Mus musculus Unknown nd A8 Brain Positive
Sample of captive micefed to meerkats at PerthZoo
A9 Mouse Mus musculus Unknown nd A9 Brain NegativeSample of captive mice fedto meerkats at Perth Zoo
A10 Mouse Mus musculus Unknown nd A10 Brain NegativeSample of captive mice fedto meerkats at Perth Zoo
A11 Mouse Mus musculus Unknown nd A11 Brain NegativeSample of captive mice fedto meerkats at Perth Zoo
B1 Kangaroo Unknown Unknown nd B1Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
B2 Kangaroo Unknown Unknown nd B2Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
B3 Pig Unknown Unknown nd B3
Porkminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
B4 Sheep Ovis aries Young nd B4
Lambminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
HK6 Kangaroo Unknown Unknown nd HK6Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
PK6 Kangaroo Unknown Unknown nd PK6Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
L2.7 Sheep Ovis aries Young nd L2.7
Lambminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
P8 Pig Unknown Unknown nd P8
Porkminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
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K1.8 Kangaroo Unknown Unknown nd K1.8Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
K2.8 Kangaroo Unknown Unknown nd K2.8
Kangaroomeatretail Positive
Meat digest of whichbradyzoites purified inPercoll
L1.11 Sheep Ovis aries Young nd L1.11
Lambminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
L2.11 Sheep Ovis aries Young nd L2.11
Lambminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
K12 Kangaroo Unknown Unknown nd K12Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
L12 Sheep Ovis aries Young nd L12
Lambminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
K13 Kangaroo Unknown Unknown nd K13Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
Mutt13 Sheep Ovis aries Adult nd Mutt13
Muttonminceretail Negative
Meat digest of whichbradyzoites purified inPercoll
HK14 Kangaroo Unknown Unknown nd HK14Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
PK14 Kangaroo Unknown Unknown nd PK14Kangaroomeat retail Negative
Meat digest of whichbradyzoites purified inPercoll
C14B Kangaroo Macropus fuliginosus AdultPositiveELISA C14 Brain Positive Wild kangaroo
C14T Kangaroo Macropus fuliginosus AdultPositiveELISA C14 Tongue Negative Wild kangaroo
C9B Kangaroo Macropus fuliginosus AdultPositiveELISA C9 Brain Positive Wild kangaroo
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C9T Kangaroo Macropus fuliginosus AdultPositiveELISA C9 Tongue Negative Wild kangaroo
J6B Kangaroo Macropus fuliginosus AdultPositiveELISA J6 Brain Positive Wild kangaroo
J6T Kangaroo Macropus fuliginosus AdultPositiveELISA J6 Tongue Negative Wild kangaroo
J10(T) Kangaroo Macropus fuliginosus AdultPositiveELISA J10 Tongue Positive Wild kangaroo
R7B Kangaroo Macropus fuliginosus AdultPositiveELISA R7 Brain Positive Wild kangaroo
R7T Kangaroo Macropus fuliginosus AdultPositiveELISA R7 Tongue Negative Wild kangaroo
Q1B Kangaroo Macropus fuliginosus AdultPositiveELISA Q1 Brain Negative Wild kangaroo
Q1T Kangaroo Macropus fuliginosus AdultPositiveELISA Q1 Tongue Positive Wild kangaroo
G21B Kangaroo Macropus fuliginosus AdultPositiveELISA G21 Brain Negative Wild kangaroo
G21T Kangaroo Macropus fuliginosus AdultPositiveELISA G21 Tongue Negative Wild kangaroo
F19B Kangaroo Macropus fuliginosus AdultPositiveELISA F19 Brain Negative Wild kangaroo
F19T Kangaroo Macropus fuliginosus AdultPositiveELISA F19 Tongue Negative Wild kangaroo
R19B Kangaroo Macropus fuliginosus AdultPositiveELISA R19 Brain Negative Wild kangaroo
R19T Kangaroo Macropus fuliginosus AdultPositiveELISA R19 Tongue Negative Wild kangaroo
H14B Kangaroo Macropus fuliginosus AdultNegativeELISA H14 Brain Negative Wild kangaroo
H14T Kangaroo Macropus fuliginosus AdultNegativeELISA H14 Tongue Negative Wild kangaroo
I14B Kangaroo Macropus fuliginosus AdultNegativeELISA I14 Brain Negative Wild kangaroo
I14T Kangaroo Macropus fuliginosus AdultNegativeELISA I14 Tongue Negative Wild kangaroo
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F8B Kangaroo Macropus fuliginosus AdultNegativeELISA F8 Brain Negative Wild kangaroo
F8T Kangaroo Macropus fuliginosus AdultNegativeELISA F8 Tongue Negative Wild kangaroo
PYC14H Kangaroo Macropus fuliginosus PYPositiveELISA PYC14 Heart Negative Pouch young of C14
PYC14B Kangaroo Macropus fuliginosus PYPositiveELISA PYC14 Brain Negative Pouch young of C14
PYC14L Kangaroo Macropus fuliginosus PYPositiveELISA PYC14 Lung Negative Pouch young of C14
PYC14Si Kangaroo Macropus fuliginosus PYPositiveELISA PYC14
SmallIntestine Negative Pouch young of C14
PYC14Li Kangaroo Macropus fuliginosus PYPositiveELISA PYC14 Liver Negative Pouch young of C14
PYC14Sp Kangaroo Macropus fuliginosus PYPositiveELISA PYC14 Spleen Negative Pouch young of C14
PYC9H Kangaroo Macropus fuliginosus PYPositiveELISA PYC9 Heart Negative Pouch young of C9
PYC9L Kangaroo Macropus fuliginosus PYPositiveELISA PYC9 Lung Negative Pouch young of C9
PYC9Si Kangaroo Macropus fuliginosus PYPositiveELISA PYC9
SmallIntestine Negative Pouch young of C9
PYC9Li Kangaroo Macropus fuliginosus PYPositiveELISA PYC9 Liver Negative Pouch young of C9
PYC9Sp Kangaroo Macropus fuliginosus PYPositiveELISA PYC9 Spleen Negative Pouch young of C9
PYJ6H Kangaroo Macropus fuliginosus PYPositiveELISA PYJ6 Heart Negative Pouch young of J6
PYJ6B Kangaroo Macropus fuliginosus PYPositiveELISA PYJ6 Brain Negative Pouch young of J6
PYJ6M Kangaroo Macropus fuliginosus PYPositiveELISA PYJ6
Skeletalmuscle Negative Pouch young of J6
PYJ6Si Kangaroo Macropus fuliginosus PYPositiveELISA PYJ6
SmallIntestine Negative Pouch young of J6
PYJ10H1 Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10 Heart Negative Pouch young of J10
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PYJ10B1 Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10 Brain Negative Pouch young of J10
PYJ10M Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10
Skeletalmuscle Negative Pouch young of J10
PYJ10Si Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10
SmallIntestine Negative Pouch young of J10
PYJ10Sp Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10 Spleen Negative Pouch young of J10
PYJ10Ki Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10 Kidney Negative Pouch young of J10
PYJ10Li Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10 Liver Negative Pouch young of J10
PYJ10Lu Kangaroo Macropus fuliginosus PYPositiveELISA PYJ10 Lung Negative Pouch young of J10
PYR7H Kangaroo Macropus fuliginosus PYPositiveELISA PYR7 Heart Negative Pouch young of R7
PYR7B Kangaroo Macropus fuliginosus PYPositiveELISA PYR7 Brain Negative Pouch young of R7
PYR7M Kangaroo Macropus fuliginosus PYPositiveELISA PYR7
Skeletalmuscle Negative Pouch young of R7
PYQ1H1 Kangaroo Macropus fuliginosus PYPositiveELISA PYQ1 Heart Negative Pouch young of Q1
PYQ1B1 Kangaroo Macropus fuliginosus PYPositiveELISA PYQ1 Brain Negative Pouch young of Q1
PYQ1M Kangaroo Macropus fuliginosus PYPositiveELISA PYQ1
Skeletalmuscle Negative Pouch young of Q1
PYG21H1 Kangaroo Macropus fuliginosus PYNegativeELISA PYG21 Heart Negative Pouch young of G21
PYG21B1 Kangaroo Macropus fuliginosus PYNegativeELISA PYG21 Brain Negative Pouch young of G21
PYG21M Kangaroo Macropus fuliginosus PYNegativeELISA PYG21
Skeletalmuscle Negative Pouch young of G21
PYF19B Kangaroo Macropus fuliginosus PYNegativeELISA PYF19 Brain Negative Pouch young of F19
PYF19H Kangaroo Macropus fuliginosus PYNegativeELISA PYF19 Heart Negative Pouch young of F20
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PYF19M Kangaroo Macropus fuliginosus PYNegativeELISA PYF19
Skeletalmuscle Negative Pouch young of F21
PYR19B Kangaroo Macropus fuliginosus PYNegativeELISA PYR19 Brain Negative Pouch young of R19
PYR19H Kangaroo Macropus fuliginosus PYNegativeELISA PYR19 Heart Positive Pouch young of R19
PYR19M Kangaroo Macropus fuliginosus PYNegativeELISA PYR19
Skeletalmuscle Negative Pouch young of R19
PY15B1B Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1 Brain Negative
Pouch young of unsampledseropositive kangaroo
PY15B1H Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1 Heart Negative
Pouch young of unsampledseropositive kangaroo
PY15B1L Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1 Lung Negative
Pouch young of unsampledseropositive kangaroo
PY15B1St Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1 Stomach Negative
Pouch young of unsampledseropositive kangaroo
PY15B1Sp Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1 Spleen Negative
Pouch young of unsampledseropositive kangaroo
PY15B1Ki Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1 Kidney Negative
Pouch young of unsampledseropositive kangaroo
PY15B1Li Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1 Liver Negative
Pouch young of unsampledseropositive kangaroo
PY15B1SkM Kangaroo Macropus fuliginosus PYPositiveELISA PY15B1
Skeletalmuscle Negative
Pouch young of unsampledseropositive kangaroo
PYH14H1 Kangaroo Macropus fuliginosus PYNegativeELISA PYH14 Heart Negative Pouch young of H14
PYH14B1 Kangaroo Macropus fuliginosus PYNegativeELISA PYH14 Brain Negative Pouch young of H14
PYH14M Kangaroo Macropus fuliginosus PYNegativeELISA PYH14
Skeletalmuscle Negative Pouch young of H14
PYI14H Kangaroo Macropus fuliginosus PYNegativeELISA PYI14 Heart Negative Pouch young of I14
PYI14B Kangaroo Macropus fuliginosus PYNegativeELISA PYI14 Brain Negative Pouch young of I14
PYR4H1 Kangaroo Macropus fuliginosus PYNegativeELISA PYR4 Heart Negative
Pouch young of unsampledseronegative kangaroo
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PYR4B1 Kangaroo Macropus fuliginosus PYNegativeELISA PYR4 Brain Negative
Pouch young of unsampledseronegative kangaroo
PYF8H1 Kangaroo Macropus fuliginosus PYNegativeELISA PYF8 Heart Negative
Pouch young of unsampledseronegative kangaroo
PYF8B1 Kangaroo Macropus fuliginosus PYNegativeELISA PYF8 Brain Negative
Pouch young of unsampledseronegative kangaroo
PYQ20H Kangaroo Macropus fuliginosus PYNegativeELISA PYQ20 Heart Negative
Pouch young of unsampledseronegative kangaroo
PYQ20B Kangaroo Macropus fuliginosus PYNegativeELISA PYQ20 Brain Negative
Pouch young of unsampledseronegative kangaroo
PY15B2B Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2 Brain Negative
Pouch young of unsampledseronegative kangaroo
PY15B2H Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2 Heart Negative
Pouch young of unsampledseronegative kangaroo
PY15B2L Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2 Lung Negative
Pouch young of unsampledseronegative kangaroo
PY15B2St Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2 Stomach Negative
Pouch young of unsampledseronegative kangaroo
PY15B2Sp Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2 Spleen Negative
Pouch young of unsampledseronegative kangaroo
PY15B2Ki Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2 Kidney Negative
Pouch young of unsampledseronegative kangaroo
PY15B2Li Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2 Liver Negative
Pouch young of unsampledseronegative kangaroo
PY15B2SkM Kangaroo Macropus fuliginosus PYNegativeELISA PY15B2
Skeletalmuscle Negative
Pouch young of unsampledseronegative kangaroo
Woylie07214B Woylie Bettongia penicillata Adult None O7214 Brain Negative Wild woylie, sudden death
Woylie07161H Woylie Bettongia penicillata Adult None O7161 Heart Negative Wild woylie, sudden death
Woylie07161B Woylie Bettongia penicillata Adult None O7161 Brain Negative Wild woylie, sudden death
PY- Pouch youngnd- not tested
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Table 5.2
Summary of polymorphisms in the B1 gene from Australian T. gondii isolates
Origin Sample 317 360 366 378 504 533 Remarks
Type I G C T G G A (Grigg and Boothroyd, 2001)
Type II/III G C C/T G C/G A (Grigg and Boothroyd, 2001)
Type X G C/G T G C A (Miller et al, 2004)
Wild kangaroo C14B G C C/T G G A/C Atypical genotype
Wild kangaroo C9B G C T A/G C/G A Atypical genotype
Kangaroo meat retail K2.8 G C T A/G C/G A Atypical genotype
Wild kangaroo R7B G C T A G A Atypical genotype
Wild kangaroo J10T G C T G C A Atypical genotype
Captive meerkat A13 G C T A C/G A Atypical genotype
Wild kangaroo J6B T G T G C A Atypical genotype
Wild kangaroo Q1T G C T G G A Type I allele
Wild kangaroo PY PYR19H G C T G G A Type I allele
Wild woylie A1b G C T G G A Type I allele
Wild woylie PY A1Ya G C T G G A Type I allele
Horse meat A7 G C T G G A Type I allele
Captive mouse A8 G C T G G A Type I allele
The numerical positions refer to the numbered sites in the published sequence (GenBankaccession no. AF179871)
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6. General discussion
6.1.Introduction
T. gondii is a significant pathogen in Australian marsupials as several outbreaks of
toxoplasmosis in captive marsupial populations have occurred and caused widespread
pathology and death (Barrows, 2006; Boorman et al., 1977; Canfield et al., 1990; Dobos-
Kovacs et al., 1974; Dubey et al., 1988; Hartley, 2006; Hartley et al., 1990; Miller et al.,
1992; Patton et al., 1986). T. gondii infection in kangaroo species is also a significant public
health concern due to the kangaroo meat trade (Holds et al., 2008). Considering the known
importance of T. gondii in Australian marsupials, it is surprising there are several gaps in
knowledge regarding the epidemiology of T. gondii in wild marsupials. This thesis aims to
tackle these gaps in knowledge by determining the prevalence of T. gondii in a range of wild
marsupial species, investigating the importance of vertical transmission and identifying the
genotype of T. gondii present in native populations. Diagnostic tools were developed and
utilised to detect T. gondii in marsupials and identify epidemiological trends. The research
undertaken has both a conservation and public health significance.
6.2.Diagnosis of T. gondii infection in Australian marsupials
A variety of tools were used in this thesis to detect T. gondii infection in marsupials. The
MAT (modified agglutination test) and ELISA (enzyme-linked immunosorbent assay) were
used to detect anti-T. gondii IgG in sera, whereas PCR (polymerase chain reaction) was
used to detect T. gondii DNA in tissue. Histology and immunohistochemistry were used to
detect T. gondii organisms and pathology in tissue.
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The MAT is the most commonly used test for serodiagnosis of T. gondii infection in
Australian marsupials (Dubey et al., 1988; Hartley and English, 2005; Lynch et al., 1993b;
Miller et al., 2003; Miller et al., 2000). The ELISA developed was found to be in very high
agreement with the MAT. The ELISA uses anti-kangaroo IgG as a secondary reagent
whereas the MAT does not utilise any species specific reagents (Ljungstrom et al., 1994). It
was found that the MAT and ELISA had a high agreement in all three macropod species
tested; the western grey kangaroo (Macropus fuliginosus), eastern grey kangaroo (Macropus
giganteus) and agile wallaby (Macropus agilis). This demonstrates the anti-kangaroo
secondary antibody utilised in the ELISA is reactive against sera from number of macropod
species. Anti-kangaroo immunoglobulin utilised in another ELISA developed to detect T.
gondii antibodies in macropods was also reactive against two different species of macropod;
the Tasmanian pademelon (Thylogale billardierii) and Bennett’s wallaby (Macropus
rufogriseus rufogriseus) (Johnson et al., 1988). Although the anti-kangaroo secondary
antibodies used in the ELISA developed were reactive against a number of macropod
species, the exact range of macropod species anti-kangaroo immunoglobulin is reactive
against is unknown. Reagent reactivity against different species may also vary between anti-
kangaroo immunoglobulin from different companies. It is recommended that for each new
macropod species tested using the ELISA developed, a number of T. gondii seropositive and
seronegative control sera from the particular species be initially screened. If the results of
the ELISA are consistent with the controls it is likely that the ELISA developed is suitable
for use in that particular macropod species.
As the ELISA developed was in high agreement with the MAT it therefore has a similar
sensitivity and specificity to the MAT. The ELISA however was more cost effective than
the commercially available MAT when large numbers of serum samples were screened. The
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ELISA may therefore be a suitable alternative to the MAT for laboratories that screen large
numbers of marsupial serum samples, such as Animal Health Laboratories (AHL) and zoos.
This ELISA could theoretically be modified to suit non-macropodine marsupials by
changing the secondary reagent and re-validating the ELISA for use.
When tissues from ELISA positive and ELISA negative western grey kangaroos were tested
using PCR for the ITS1 sequence (Nandra and Grigg, manuscript in preparation) absolute
agreement was observed. Out of 9 seropositive adult kangaroos tested using PCR, nested
PCR for the ITS1 sequence was positive for all 9 animals. However, nested PCR for the B1
gene was only positive in 6 out of 9 seropositive adult animals. Non-nested PCR for the B1
gene was positive in 1 out of 9 seropositive adult animals. The results indicate that the
nested PCR used for the ITS1 sequence (Nandra and Grigg, manuscript in preparation) has a
comparable sensitivity to the T. gondii ELISA developed. The nested PCR for the B1 gene
used (Grigg and Boothroyd, 2001) showed a lower sensitivity than the nested PCR for the
ITS1 sequence. This may be readily explained by the difference in copy number between
the different loci amplified; the ITS1 sequence is present in 110 copies in the T. gondii
genome compared to a copy number of 35 for the B1 gene (Hurtado et al., 2001). Therefore
it is expected that PCR targeting a higher copy number sequence is more sensitive at
detecting T. gondii DNA (Switaj et al., 2005). Non-nested PCR for the B1 gene detected T.
gondii DNA in fewer samples than nested PCR for the B1 gene. This supports results of
Pujol-Rique et al (1999) which demonstrate that nested PCR is more sensitive than non-
nested PCR for detecting T. gondii DNA in clinical samples. (Pujol-Rique et al., 1999)
DNA degradation may also have influence on PCR sensitivity (Lahiri and Schnabel, 1993).
Non nested PCR detected T. gondii DNA in two DNA samples that were negative using a
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nested ITS1 PCR and nested B1 PCR. This result may be explained by the time when which
DNA samples were tested for T. gondii using different primers. Non nested PCR for the B1
gene was the first PCR to be optimised and validated during the period of this study.
Similarly, non-nested B1 gene primers were the first to be utilised against DNA samples
extracted from tissue. Nested ITS1 and nested B1 PCR primers were applied to DNA
samples after a period of delay, and many freeze-thaw cycles. DNA is degraded by long
term storage and freeze-thaw cycles (Lahiri and Schnabel, 1993), and this may explain why
some DNA samples that were initially positive using non-nested B1 primers were not
positive using other primers. It is highly unlikely that DNA samples that were positive by
the non-nested PCR for the B1 gene and negative by other primers were false positive
results. This is because DNA extraction water controls and PCR water controls were both
negative for T. gondii when the DNA samples in question were PCR positive.
No T. gondii organisms were observed using histology or immunohistochemistry in this
study, even in tissues that were PCR positive. Two western grey kangaroo pouch young, one
adult woylie (Bettongia penicillata) and one meerkat (Suricata suricatta) were positive for
T. gondii using PCR; however no T. gondii organisms were observed in tissues of these
animals using histology or immunohistochemistry. Upon histology of a mammary gland that
was T. gondii PCR positive in an adult woylie, a focal area of inflammation without
tachyzoites or bradyzoites was observed. No inflammation was observed in other tissues
examined from the woylie, including heart, brain and skeletal muscle. These tissues were
also PCR negative. A similar situation to that observed in the woylie was seen in a PCR
positive meerkat tested. A meerkat with a PCR positive brain sample had
meningoencephalitis, with no visible tachyzoites. Inflammation without the presence of
visible tachyzoites is described in the literature in some cases of toxoplasmosis (Canfield et
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al., 1990). In addition upon experimental T. gondii infection in immunocompetent mice, the
number of tachyzoites peaked at approximately 12 days. By 21 days it was difficult to
identify tachyzoites in any organ, even when immunocytochemistry was used (Ferguson and
Dubremetz, 2007). The combination of PCR positive brain samples and
meningoencephalitis in the meerkat examined strongly suggests that the
meningoencephalitis observed was due to T. gondii infection. T. gondii associated
meningoencephalitis has previously been reported in meerkats (Juan-Sallés C et al., 1997)
and is reported in other animals including Californian sea otters (Miller et al., 2004),
dolphins (Dubey et al., 2003e; Jardine and Dubey, 2002), wallabies (Basso et al., 2007) and
a wombat (Skerratt et al., 1997).
One limitation of the histological analysis of western grey kangaroo pouch young is that
brain samples were not examined. This is because the young were killed by blunt trauma to
the head and the cranial cavity damage induced at death caused postmortem autolysis in the
immature brain tissue. Brain samples were still used for PCR, however none were positive
for T. gondii DNA. The only positive tissue samples in both PCR positive western grey
kangaroo pouch young was heart muscle. Despite this no inflammation or tachyzoites were
observed in the PCR positive heart muscle using histology or immunohistochemistry. A
similar situation occurs in chronically infected tammar wallabies (Reddacliff et al., 1993). In
two chronically infected tammar wallabies, minimal inflammation and no tachyzoites were
observed on histology. Due to the exclusion of brain tissue from histological analysis it
cannot be proven that there was no T. gondii associated pathology in the two PCR positive
western grey kangaroo pouch young; however, the observation of no inflammation in all
organs examined strongly suggests these animals were not suffering clinical toxoplasmosis.
This is because clinical toxoplasmosis is documented as causing inflammation in multiple
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organs in the large majority of marsupials screened histologically (Canfield et al., 1990;
Obendorf and Munday, 1983, 1990; Patton et al., 1986; Reddacliff et al., 1993).
6.3.Epidemiology of T. gondii in Australian marsupials
A T. gondii seroprevalence of 15.5% (95%CI: 10.7-20.3) was observed in adult western
grey kangaroos in the Perth metropolitan area using an ELISA. A similar seroprevalence
level of 17.7% (Johnson et al., 1988) and 15.5% (Turni and Smales, 2001) was found in
Tasmanian pademelons and bridled nailtail wallabies (Onychogalea fraenata) respectively.
The observation of chronically infected, live macropod marsupials in the wild is in contrast
to the observation of toxoplasmosis causing pathology and death in several collections of
captive macropods (Boorman et al., 1977; Canfield et al., 1990; Dobos-Kovacs et al., 1974;
Dubey et al., 1988; Miller et al., 1992; Patton et al., 1986). The result of seropositive
western grey kangaroos surviving in the wild demonstrates that T. gondii infection does not
always cause severe death and pathology in marsupials and that a secondary factor may be
needed to induce clinical toxoplasmosis. It is thought that subclinical, chronic T. gondii
infection in marsupials can become acute disease by exposure to secondary factors such as
capture, transportation, captivity, malnourishment and extreme weather (Arundel et al.,
1977; Beveridge, 1993; Obendorf and Munday, 1983, 1990). T. gondii genotype may also
have an effect on the clinical manifestation of T. gondii in Australian marsupials. Six out of
the 34 seropositive western grey kangaroos in the seroprevalence study had T. gondii DNA
sequenced. Five out of the six kangaroos tested harboured atypical T. gondii genotypes, and
it is possible that these T. gondii genotypes are less virulent to kangaroos than other T.
gondii genotypes that have previously infected kangaroos in captivity and caused
toxoplasmosis.
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Intermediate hosts of T. gondii may acquire infection in one of three ways; the ingestion of
viable T. gondii oocysts shed in felid faeces, the ingestion of viable T. gondii bradyzoites in
tissue cysts or vertical transmission (Obendorf and Munday, 1990). The results of
seroprevalence studies in chapter 3 provide epidemiological evidence that oocysts play an
important role in the transmission of T. gondii in Australian marsupials. Results of vertical
transmission studies in chapter 4 indicate that vertical transmission occurs in marsupials. In
addition, the discovery of atypical strains of T. gondii in Australian marsupials in chapter 5
leads to questions regarding the origin and transmission of these atypical strains. Overall,
results suggest that oocyst transmission is important in maintaining T. gondii marsupial
populations and that vertical transmission also occurs and may contribute to the prevalence
of T. gondii marsupial populations.
Seroprevalence studies of western grey kangaroos in Perth, WA found that the
seroprevalence of T. gondii in males was significantly less than in females (p=0.038). One
difference between male and female western grey kangaroos that could explain their
difference in exposure to T. gondii within the same environment is their difference in
feeding habits. Females are able to graze closer to the ground than male kangaroos,
particularly when there is a scarcity of graze (Newsome, 1980). Other differences in
behaviour between male and female kangaroos that may influence the level of exposure to
T. gondii are not known. The ability of female kangaroos to graze closer to the ground
however can explain the significant difference in seroprevalence between male and female
western grey kangaroos, as grazing closer to the ground increases exposure to oocysts in
soil. Therefore females with a greater exposure to T. gondii oocysts in soil would be
expected to have a higher seroprevalence of T. gondii than their male counterparts. An
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alternatitive reason why females had a significantly higher T. gondii seroprevalence than
males is recrudescence of T. gondii during pregnancy/lactation. A rise in anti-Neospora
caninum antibodies is known to occur during pregnancy in cattle, and may be associated
with recrudescence of N. caninum infection and vertical transmission (Conrad et al., 1993;
Haddad et al., 2005; Pare et al., 1997; Wouda et al., 1999). It has not yet been proven that
recrudescence of T. gondii may occur during pregnancy/lactation in marsupials. However,
results of vertical transmission studies in chapter 4 suggest vertical transmission can occur
during chronic T. gondii infection and this may be associated with recrudescence of T.
gondii in female marsupials. Further studies in marsupials need to be undertaken to identify
the mechanism by which vertical transmission of T. gondii occurs and confirm if
recrudescence of T. gondii in pregnant/lactating marsupials takes place.
A low combined seroprevalence of 0.625% was observed in marsupials located in areas free
from felids. In comparison a moderate combined seroprevalence of 8.31% was observed in
marsupials located in areas where felids may roam. A case control study undertaken found a
statistically significant difference in seroprevalence between marsupials located in areas
where cats may roam and marsupials located in areas without cats. It was calculated that
marsupials located in areas where cats may roam are 14.2 times more likely to be T. gondii
seropositive. Felids are the only known animals capable of shedding oocysts and it is
expected felid exposure mirrors oocyst exposure. The results of this seroprevalence study
enforce the results of previous epidemiological studies in pigs (Dubey et al., 1997a),
humans in Costa Rica (Frenkel and Ruiz, 1981) and sheep (Munday, 1972) that indicate
felids are important in the transmission of T. gondii. The study undertaken in marsupials is
the first of its kind in demonstrating the significance of felids on T. gondii prevalence in
wild marsupials.
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Unlike oocyst transmission, which is a well known form of T. gondii transmission, little is
known regarding the role of vertical transmission in the maintenance T. gondii infection in
animal and human populations. Vertical transmission is often thought of in terms of its
ability to cause disease in newborns rather than in terms of its effect on T. gondii prevalence
(Marshall et al., 2004). A study was undertaken to determine the occurrence of vertical
transmission of T. gondii in Australian marsupial species. A number of tests were
undertaken in marsupial dams that were infected with T. gondii and their offspring.
Evidence of vertical transmission was found in two marsupial species tested; western grey
kangaroos and woylies. The offspring of 10 seropositive western grey kangaroos were tested
for T. gondii infection using PCR, comparative immunoblots, histology and
immunohistochemistry. Of the 10 pouch young tested, each from different dams, two were
positive for T. gondii infection using PCR. Negative DNA extraction and PCR controls from
pouch young of seronegative dams remained PCR negative. PCR positivity had a high
correlation with adult seropositivity, as all seropositive dams tested using PCR were PCR
positive and all seronegative dams tested using PCR were PCR negative. This indicates the
two PCR positive western grey kangaroo pouch young were infected with T. gondii and the
results were not false positive. Histology and immunohistochemistry found no evidence of
T. gondii associated pathology in either of the two PCR positive pouch young. This suggests
neither of the positive pouch young had clinical toxoplasmosis. Furthermore, none of the
seropositive western grey kangaroo dams were positive for IgM, which indicates the dams
were all chronically infected with T. gondii when they and their offspring were sampled.
The results indicate that chronically infected dams can transmit T. gondii vertically and T.
gondii infected offspring can remain healthy, which supports the hypothesis that vertical
transmission is capable of maintaining T. gondii infection in wild marsupial populations.
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Vertical transmission of T. gondii was also demonstrated in a woylie dam and its pouch
young. The mammary gland of a woylie dam was PCR positive and the brain of its
corresponding young, which was unfurred and had never left the pouch, was also PCR
positive. It is unknown if the woylie dam was acutely infected with T. gondii as no sera was
available for testing in the woylie. In addition, it is unknown if the woylie pouch young was
clinically affected with toxoplasmosis as no tissue samples were available for histology.
However, the unique result of a mammary gland of a woylie dam and the brain of its
unfurred pouch young being PCR positive is suggestive of milk transmission of T. gondii
from the woylie dam to the pouch young via the mammary gland. Results suggest that when
vertical transmission of T. gondii occurs in marsupials, it may occur via the mammary
gland. This form of vertical transmission is expected in marsupials because marsupial young
are born at a very immature state and milk is the source of sustenance which enables young
to develop to a stage where they can leave the pouch (Dawson, 1995; Tyndale-Biscoe and
Renfree, 1987). If marsupial young were infected in utero, their immature state would likely
cause them to succumb to toxoplasmosis and die (Dubey et al., 1988).
Sequencing of B1 PCR products of DNA from Australian tissue samples found 7 out of 13
samples sequenced to have an atypical genotype. This was the first study to molecularly
characterise T. gondii DNA from Australia. The 7 T. gondii DNA isolates had SNPs in the
B1 gene that were different from any strains documented to date. Six unique genotypes were
identified out of the 7 atypical isolates; two out of the 7 isolates had the same unique
sequence at the B1 gene whereas the other 5 isolates each had different combinations of
SNPs at the B1 gene. The majority of the T. gondii DNA isolates sequenced were from
Australian wildlife. Out of the 13 isolates sequenced, 8 were from kangaroos and 2 were
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from woylies. In addition isolates from a meerkat, mouse and horse were sequenced. The
discovery of atypical strains of T. gondii in Australia leads to further questions regarding the
origin and transmission of these atypical strains.
Further loci need to be examined to ascertain if the atypical isolates found are recombinant
or novel strains. If some isolates are novel, which is feasible considering many of the
isolates found have unique SNPs, a worthy question is; how and when were these novel
strains introduced to Australia? A study by Su et al (2003) suggests that novel (exotic)
strains were derived from genetic crosses of ancestral lineages more than 10000 years ago.
Clonal lineages, which predominate in Europe and North America, were analysed as arising
from a more recent genetic cross to novel strains (Su et al., 2003). If the T. gondii isolates in
marsupials are found to be novel strains unique to Australia and the analyses of Su et al
(2003) hold true, this would suggest that novel T. gondii strains were in Australia long
before European settlement and the introduction of felids. If T. gondii was present in
Australia before the introduction of felids, the only known way T. gondii may be have been
transmitted is via tissue cyst transmission and vertical transmission. Another possibility is
that some now extinct non-felid Australian native species had the ability to serve as a
definitive host.
Suggestions from Ajzenberg et al (2004) regarding the origin of novel and recombinant
strains are different from Sue et al (2003). Ajzenberg et al (2004) suggests recombinant and
novel strains arise in wild populations with a high host diversity, where enhanced sexual
propagation in felid hosts causes genetic recombination (Ajzenberg et al., 2004). Many
believe that the clonal strains of T. gondii are those which have successfully adapted to
domestic hosts (Ajzenberg et al., 2004; Lehmann et al., 2003; Su et al., 2003). Ajzenberg et
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al (2004) suggests that clonal strains of T. gondii predominate in domestic animals and
humans whereas recombinant and novel strains flourish in ecologically rich wild
environments. There is speculation that the predominance of clonal strains in North America
and Europe is due to their production of a small range of domestic meat-producing animals
and the subsequent propagation of clonal strains via tissue cysts (Ajzenberg et al., 2004;
Darde et al., 2007; Dardé et al., 2008). Seroprevalence results from chapter 3 demonstrate
that oocyst transmission (through felid hosts) is common in wild marsupials; this high rate
of oocyst transmission may theoretically result in increased sexual recombination in felid
hosts and the development of non-clonal T. gondii genotypes in wild marsupial populations
in Australia. To better understand the transmission of atypical strains in Australia, more T.
gondii DNA isolates need to be obtained from domestic Australian hosts, including
domestic felids. An in depth study comparing wildlife and domestic isolates from Australia
to those found in the rest of the world would provide important information regarding the
origin and transmission of T. gondii in Australia.
Out of 10 tissue samples from wild Australian marsupials that were PCR positive at the B1
gene, 6 had atypical genotypes and 4 had a type I allele. This preliminary result suggests a
high percentage of wild Australian marsupials are infected with atypical or type I strains of
T. gondii. It is unknown if strain type affects disease severity in Australian marsupials. This
is a worthy area for future studies as it may explain why some marsupials display more
severe disease than others when infected with T. gondii. Different strains have different
levels of virulence in mice, with type I strains having an LD100 of one parasite and type II
and III strains having an of LD100 of several thousand parasites (Boothroyd and Grigg,
2002). In addition, type I strains are associated with severe ocular toxoplasmosis in
immunocompetent humans (Boothroyd and Grigg, 2002; Grigg et al., 2001b; Vallochi et al.,
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2005). Atypical type X strains have also been associated with severe meningoencephalitis in
Californian sea otters (Miller et al., 2004).
Previous experimental infections of marsupials with T. gondii may give clues as to whether
different strains cause different diseases in marsupials. The S48 strain of T. gondii, which is
mouse virulent (Fazaeli et al., 2000; Innes and Mattsson, 2007; Wilkins et al., 1988) caused
death in 4 out of 4 seronegative tammar wallabies experimentally infected (Lynch et al.,
1993a). Similarly, P89/VEG strain which is recombinant type I/III and SAG1 type I (Howe
and Sibley, 1995; Mondragon et al., 1998b) caused death in 4 out of 4 experimentally
infected eastern barred bandicoots (Bettiol et al., 2000a; Bettiol et al., 2000b). Conversely
the mouse avirulent type II ME49 strain (Ferreira et al., 2006) did not cause 100% mortality
in tammar wallabies and caused death in 7 out of 9 wallabies infected (Reddacliff et al.,
1993). In addition, the mouse avirulent pork I strain (Johnson, 1988) caused 0% mortality in
three experimentally infected eastern grey kangaroos (Johnson et al., 1989). The differing
mortality rates seen in marsupials experimentally infected with different strains of T. gondii
suggest that mouse virulent (or SAG1 type I) strains are more virulent in marsupials than
mouse avirulent strains. It must be noted that certain species of marsupial may be more
susceptible to death by toxoplasmosis than others and this may have affected the differing
responses to experimental infections with different strains. In addition, the response of
marsupials to experimental infection in captivity compared to natural infection in the wild
may be very different. Further studies need to be undertaken to analyse the genotype of T.
gondii found in different species of naturally infected marsupials, and the clinical
signs/pathology associated with infection with different T. gondii genotypes. This
information would be valuable in linking T. gondii genotype to T. gondii disease
manifestation in Australian marsupials.
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6.4.Suggestions for future research
Despite the importance of T. gondii infection in Australian marsupials, there is no
serological test that has been assessed for sensitivity and specificity in marsupials. Even the
sensitivity and specificity of the MAT, which is the most commonly used test to detect anti-
T. gondii IgG in marsupials, has not been determined in marsupials. Some studies have
observed a high correlation between MAT positivity and chronic T. gondii infection (Dubey
et al., 1988; Johnson et al., 1989; Obendorf et al., 1996). However, the sensitivity and
specificity of the MAT needs to be determined, for a number of marsupial species, to better
understand results of marsupial T. gondii seroprevalence studies. The sensitivity of the
MAT would affect the amount of false negatives in any given population. Likewise, the
specificity of the MAT would affect the number of false positives. It is unknown if
antibodies against other coccidia infecting marsupials may cross react with MAT reagents
and cause false positive results. To determine the sensitivity and specificity of the MAT in
marsupials, it would be ideal to undertake experimental T. gondii infections in a number of
marsupial species, with each marsupial species representative of one marsupial genus.
Marsupials may be tested using the MAT before and after experimental infection with T.
gondii. In addition, MAT negative marsupials may be experimentally infected with other
coccidia such as a Sarcocystis species and Neospora species to determine if cross reactivity
occurs. The probable reason why such studies have not yet been undertaken is that costs of
such a study may outweigh the benefits. Experimental infection of marsupials with T. gondii
often results in high mortality rates (Bettiol et al., 2000a; Bettiol et al., 2000b; Lynch et al.,
1993a; Reddacliff et al., 1993), sometimes before IgG can be detected (Lynch et al., 1993a).
The costs associated with killing a large number of marsupials may outweigh the benefits of
136
determining the sensitivity and specificity of the MAT. An alternative to experimental
infection is to obtain paired sera and tissue samples from dead marsupials and screen tissues
for T. gondii infection using bioassay and PCR. Sera could then be tested using the MAT
and the MAT results correlated with the bioassay and PCR results. One cost of this method
is that it may take long periods of time to obtain a sufficient amount of paired sera and
tissue samples, particularly from rarer marsupial species. Such a study may be done in
collaboration with zoos, wildlife parks, veterinary clinics and Animal Health Laboratories
(AHL) in Australia. The screening of marsupial tissue for T. gondii using bioassay and PCR
as mentioned above would also provide a better understanding of the range of T. gondii
strains present in Australian marsupials. T. gondii DNA detected using PCR could be
sequenced to determine the genotype of T. gondii infecting marsupials. As mentioned in
section 7.3, it would be ideal to correlate T. gondii genotype with clinical signs and
pathology.
Comparison of the T. gondii strains present in Australian marsupials to T. gondii strains
present in domestic hosts in Australia should be undertaken, as mentioned in section 7.3. It
would provide important information regarding the origin and transmission of T. gondii in
Australia. If the atypical strains of T. gondii found in marsupials cannot be found in felids,
this would suggest the atypical strain cannot be transmitted by cats and therefore point to
other felid-free methods of transmission. Felid-free methods of T. gondii transmission
include vertical transmission and tissue cyst transmission. It may also be possible that non-
felid Australian native species have the ability to shed T. gondii oocysts. Alternatively, if it
is found that domestic hosts have a significantly higher rate of carrying clonal T. gondii
strains than wild hosts, this may suggest the presence of a sylvatic cycle of T. gondii
transmission.
137
Additional research into the frequency of vertical transmission in Australian marsupials is
also warranted. Vertical transmission of T. gondii in chronically infected marsupials was
demonstrated in this study. The results emphasise the need for further studies using a larger
number of marsupials to determine the frequency of vertical transmission in range of
marsupial species. The ability of T. gondii to be vertically transmitted through successive
generations is also a matter of interest. If T. gondii can be transmitted through successive
generations in marsupials, this would imply vertical transmission is a major method of
maintaining T. gondii infection in marsupial populations. The main restriction of testing if
T. gondii can be transmitted through successive generations is that marsupials of the first
generation should be kept in a T. gondii free environment while conceiving the second
generation. Alternatively, T. gondii seronegative marsupials could be used as negative
controls for environmental contamination where the experiment is held.
6.5.Concluding remarks
Results show that the ELISA developed may be a suitable cost effective alternative to the
MAT. T. gondii is present in western grey kangaroo, eastern grey kangaroo and woylie
populations in Australia. Marsupials located in areas were cats may roam are more likely to
be T. gondii seropositive than marsupials located in areas without cats. In addition, vertical
transmission occurs in marsupials, possibly via the mammary gland. Marsupial vertical
transmission studies also suggest vertical transmission can occur during chronic infection
and result in clinically unaffected offspring. However, further studies analysing the
frequency of vertical transmission and the occurrence of vertical transmission in successive
generations needs to be undertaken to better understand the importance of vertical
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transmission in maintaining T. gondii infection in marsupial populations. Atypical T. gondii
genotypes were found in Australian marsupials in this thesis, and this leads to several
further questions regarding the origin, transmission and virulence of these atypical
genotypes.
139
References
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