Associations between gastrointestinal

parasites and Nidovirus infection in

Western Australian

Shingleback Lizards (Tiliqua rugosa)

by

Jazmin Lilibeth Martinez

Bachelor of Science

This thesis is presented for examination for the Bachelor of Science Honours in

Conservation Wildlife Biology.

School of Veterinary and Life Sciences

Murdoch University, Perth

2019

iii

Author’s Declaration

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.

Jazmin Lilibeth Martinez

v

Abstract

Australia’s native reptile, the Shingleback lizard (Tiliqua rugosa) is under threat from a variety

of causes including a newly discovered Nidovirus(1). This flu has caused an influx of

Shinglebacks into rehabilitation centres, demonstrating upper respiratory tract infections

(URTI). Therefore, understanding other factors that can impact on the progression of this

infection is extremely important for the conservation of the Shingleback lizard population. Such

factors include the effects of co-infections, which are composed of two or more pathogens

working synergistically or antagonistically against each other. Interactions seen in co-infections

tend to cause enhancement of pathogenesis(3). In this study we investigated the association

between gastro-intestinal (GI) parasites and the Nidovirus within the Tiliqua rugosa population.

The main aims and hypothesis of this study is to investigate if any association exists between

the abundance and diversity of individual parasitic loads, with the presence/absence of the novel

Tiliqua rugosa Nidovirus. This will be completed by three objectives, (i) obtaining baseline data

on the prevalence of infection, (ii) confirming the proportion of animals infected by coinfections

and (iii) examine any possible relationship seen in co-infected Shingleback lizards.

Samples were collected from Shingleback lizards entering Native ARC Rehabilitation Centre

and Kanyana Wildlife Rehabilitation Centre. Oral swabs were taken for virology analysis which

was completed through polymerase chain reaction (PCR) technology. Faecal samples were

collected when naturally produced and were analysed at Murdoch University laboratories,

through malachite stained faecal smear(4) and zinc sulphate faecal flotation techniques(5).

There were three main findings from this study. First, prevalence of infection was relatively

high, with 73 % of Shinglebacks infected with at least one species of gastro-intestinal parasite

and 58.1% of Shinglebacks infected with the T. rugosa Nidovirus. Secondly, co-infections with

gastro-intestinal parasites and Nidovirus were common; 48.4% of Shinglebacks carried both

parasitic and viral infections, while 26% had only parasitic infections and 9% had only viral

infections. Finally, evidence was found corroborating my initial hypothesis that the major

vi

clinical signs of viral infection, discharge from the eyes or nose, were enhanced in Shinglebacks

which were co-infected with the virus and gastro-intestinal parasites.

This study suggests that the Nidovirus seen in the Tiliqua rugosa population is part of the multi

dynamic route of disease seen in reptiles (6), in particular an emerging pattern between co

infection with GI parasites suggest synergistic interaction may be a possible factor in the

progression of pathogenesis in this viral infection. Therefore, possible immunological

enhancement of the virus or/and the parasites may be seen within an individual host. It is not

clear yet if this enhancement of infection is caused due to the immunosuppressive

characteristics of the virus or due to high parasitic abundance causing immunological stress on

the hosts.

This is the first comprehensive study of gastrointestinal parasitism in wild populations of

Tiliqua rugosa and the first study of co-infections of gastro-intestinal parasites and Nidovirus.

This paper will allow a clearer understanding of natural co-infections, synergistic and/or

antagonistic relations between pathogens and the potential cause of pathogenesis.

vii

Acknowledgements

I would like to thank Native ARC Rehabilitation centre for their amazing efforts during this

study, in particular Veterinarian treatment team Dr Szou Whua Bosci and Dr Meg Rodgers who

made the time to be involved with this study even with time constraints from being busy with

other Native ARC procedures. Their ability to fit me in during such busy periods was greatly

appreciated and their team of volunteers were of much aid in the collection of samples during

the entire study period. I would like to thank both Dr Szou and Dr Meg for sending over

information of the Shingleback faecal analysis which was not performed by Murdoch, this

assistance plus their willingness to aid in anyway was kindly appreciated. Lastly Dean Huxley,

manager at Native ARC, thank you kindly for being so welcoming of me and allowing the study

to progress even with the busy schedule Native ARC had to upkeep.

A massive thank you goes to the Kanyana Wildlife Rehabilitation Centre, in particular Tasha

Hennings (Hospital Manager) who was an absolute pleasure to work with. The entire volunteer

team at Kanyana aided in every aspect of this studies sample collection, they were extremely

professional, and kind hearted throughout my entire time with them. This study owes a great

deal to the constant aid of Kanyana and their excellent service. There are a few volunteers who

must be acknowledged for that involvement in parasitic analysis of faecal samples which were

unable to be analysed at Murdoch. These Lovely volunteers are G. Thomas, N. Jardine, M.

Pryor and B. Brice. Carol Jackson was one of the volunteers who’s aid in this project was

extremely important, not only did she provide faecal analysis of Shinglebacks she also

generously provided all pictures of the Shinglebacks symptoms for this paper and assisted in

any way possible.

Further acknowledgment is extended to Murdoch and the Murdoch Parasitology team, with their

expert advice this study was able to progress very smoothly. I would personally like to thank Dr

Amanda Ash head supervisor of this study, Dr Alan Lymbery co-supervisor and Dr Mark

O’Dea co-supervisor. Their help every day was extremely useful and encouraging, their expert

advice, opinions and guidance were of much aid in this study. Amanda’s extensive knowledge

viii

of parasitic species provided a great depth of knowledge for me to learn from, Alan’s expert

opinion from years of academia aided greatly into shaping this study into a coherent paper,

lastly Mark’s knowledge and previous work on the Tiliqua Rugosa study provided the core to

this study and without his expert knowledge on virology this paper will not exist. I owe a great

deal to these three wonderful professionals and cannot thank them enough for their life lessons,

academic excellence and personal guidance during this entire experience. In particular I am

extremely grateful of the time and effort they placed into me so that I can further improve this

paper, there selflessness and generosity will never be forgotten.

I would like to thank my family, mostly my parents Yussi and Diego Martinez for whom which

I would not have made it passed my undergraduate stage. I would like to thank them for always

encouraging me to keep going and supporting me during many lows in my academic years. A

big thank you is extended to my partner, Jamie Bozman who tolerated many emotional days of

worry revolving around this paper and always supporting me to keep studying and furthering

my degree. I would also like thank my brother Anthony Martinez, who stayed with me at

university till 3 am many nights to encourage me to keep going. His support means the world to

me.

Great appreciation goes towards Monique Smith fellow Honours Graduate whom without my

statistical analysis would not exists, her knowledge and experience with R studios was beyond

outstanding and her patients with me was greatly appreciated, I owe my entire result section to

Monique and cannot thank her enough for her help. Finally, I would like to thank my dear

friends Paris Brown and Brittny Forsyth who kept me smiling during writing this paper and

always encouraged me to keep going.

ix

Table of Contents

Declaration ........................................................................................... iii

Abstract ................................................................................................ v

Acknowledgments ............................................................................ vii

List of figures .................................................................................... xi

List of Tables .................................................................................. xiii

List of Abbreviations ...................................................................... xiii

Chapter 1 ............................................................................ Introduction 15

1.1 ..................................................................................... Co-infection 15

1.2 ........................................................... Host response to co-infection 15

1.3 .......................Synergistic and antagonistic impacts of co-infection 16

1.4 ......................................................... Parasitic infections in wildlife 18

1.5 .................................................................................. Tiliqua rugosa 19

1.6 .............................................. Parasitic infections in Tiliqua rugosa 20

1.6.1 ..................................................................................... Protozoans 21

1.6.2 ...................................................................................... Helminths 24

1.6.3 ..............................................................................Viral infections 24

1.6.3.1 ................................................................................. Nidovirales 24

1.6.3.2 ............................................................................ Coronoviridae 25

1.6.3.3 ............. Pathogenesis of the novel Nidovirus in Tiliqua rugosa 28

1.7 ....................................................................... Aims and Hypothesis 30

Chapter 2 .......................................................................... Methodology 31

2.1 ............................................................................. Sample collection 31

2.2 ................................................ Measurement of age and body mass 32

2.3 ...................................................... Measurement of body condition 34

2.4 .............................................. Measurement of viral symptom score 34

2.5 ................................................................................ Faecal Analysis 34

2.5.1 ................................................................................. Faecal Smear 35

2.5.2 ............................................................................. Faecal Flotation 35

2.5.3 ........................................................................................ Analysis 35

2.6 ........................................................................................... Virology 36

2.7 ............................................................................ Statistical analysis 36

x

Chapter 3 .................................................................................... Results 37

3.1 ............................................................. Shinglebacks demographics 37

3.2 ......................................................................................Parasitology 39

3.3 ........................................................................................... Virology 43

3.4 ..................................................................................... Co-infection 44

3.5 ........................................................ Clinical signs of viral infection 46

Chapter 4 ................................................................. General discussion 49

4.1 ..................................... Prevalence of parasitic and viral infections 49

4.2 ..................................... Co-infections with parasites and Nidovirus 52

4.3 ............................................... Co-infections and viral clinical signs 53

4.4 ............................................. Study limitations and further research 56

Chapter 5 .............................................................................. References 57

Chapter 6 ................................................................................ Appendix 62

xi

List of Figures

Figure 1.1 Over of potential consequences of multiple infections with regards to hosts manipulation (2). 16

Figure 1.2 Maximum likelihood phylogenetic tree of Nidovirus Tiliqua rugosa (1) 27

Figure 1.3. a= healthy Shinglebacks body condition, b= mild URTI body condition, c= severe URTI body

condition. Picture provided by Tasha Henning’s, hospital manager,

Kanyana Wildlife Rehabilitation Centre. 29

Figure 2.1. a= eye discharge, b= mucous in oral cavity. Picture provided by Tasha Henning,

Kanyana Wildlife Rehabilitation Centre. 34

Figure3.1. Distribution of the original locations in which Shinglebacks lizards were found before entering

rehabilitation centres. Dot size indicates number of animals collected at a location. 37

Figure 3.2 Relationship between snout-vent-length and weight in the sampled Shinglebacks. Regression line 38

Figure 3.3 a-h. Parasitic species found during faecal analysis 39

Figure 3.4 Prevalence of parasite taxa in adult and juvenile Shinglebacks 41

Figure 3.5 Mean number of parasites/individual host in adult and juvenile Shinglebacks. 42

Figure 3.6 Prevalence of Nidovirus l infection in adult and juvenile Shinglebacks. 43

Figure 3.7 Prevalence (%) of viral and parasitic infections in Shinglebacks 44

Figure 3.8. Percentage Prevalence of Viral Infection in Accordance to Parasitic Infection 44

Figure 3.9 Mean number of parasite taxa in Shinglebacks with and without Nidovirus infections 45

Figure 3.10 Percentage prevalence of viral infection in accordance to viral symptom score 46

Figure 3.11 Mean body condition score of Shinglebacks with and without Nidovirus infection. 47

Figure 3.12. Percentage Prevalence of Viral Symptoms in Accordance to Parasitic Infection 48

Figure 3.13 Mean number of parasite taxa for virally infected Shinglebacks that showed or didn’t show clinical

signs 48

Figure 6.1 White gum commonly seen in shinglebacks infected with URTI infection 62

Figure 6.2 Thickened eyed discharge 62

xii

List of Tables

Table 1.1 Protozoans found in the Tiliqua rugosa population 22

Table 1.2 Helminths found in the Tiliqua rugosa 23

Table 2.1. Age and body mass estimation per the weight and length of the Shinglebacks. Information access

provided by Tasha Hennings, hospital manager, Kanyana Wildlife Rehabilitation 32

Table 3.1 Prevalence (%) and mean intensity of parasite taxa found in sampled Shinglebacks 40

Table 3.2 Mean intensity parasite taxa in adult and juvenile Shinglebacks 42

xiii

List of Abbreviations

PI- Prime investigator

IM – Intra muscular

PO- Per oral

URTI- Upper respiratory tract infection

URT- Upper respiratory tract

PCR Polymerase chain reaction

RT-PCR- Reverse real time polymerase chain reaction

RNA- Ribonucleic acid

HIV- Human immunodeficiency viruses

DWV- Deformed wing virus

15

Chapter 1 Introduction

1.1 Co-infection

The term co-infection refers to the co-existence of multiple infections agents in the same hosts.

Co-infection includes other nomenclature such as, poly parasitism, mixed infection, secondary

infection or concurrent infection; (7) refers to the co-existence of multiple infectious agents in

the same host. Individuals can be subjected to a wide variety of pathogens including viruses,

bacteria, fungi, yeast, protozoa, and eukaryotic parasites (8). These pathogens can be found in

homologous (same species) or heterologous (different species) relationships within the host,

with antagonistic or synergistic effects on each other (9).

Co-infections are a major problem for human and animal health (10). The interactions seen in

co-infections are complex, therefore there is a lack of understanding for disease progression and

dynamics in co-infected hosts (11). Co-infections make up 30% of infectious diseases in first

world countries and up to 80% in developing countries. Co-infections are also ubiquitous in

both domestic animals and wildlife (12) . Research in co-infection patterns in wildlife diseases

is extremely important for understanding the effects it has on the natural ecosystem and on

populations of wild animals, but also in the advancement of animal disease and zoonotic

treatment. Due to the uniquely variable nature of co-infections studying the relationships can be

difficult, especially in natural ecosystems (3, 7, 8).

1.2 Host response to co-infection

One angle of investigation is looking at the interactions of pathogens and the hosts, these

interactions include immune response and pathogen dynamics. Exposure to pathogens allows

for the progression of immune memory and enhancement of host-fitness through evolutionary

change (11-13). The presence of foreign antigens initiates a specific protective immune

response, which leads to a series of cellular and humoral events within the body. The host’s

immune response directs itself towards the humoral or cellular response depending on the

antigen present. This process may be altered by pathogen dynamics which may favour stabilized

pathogenic communities or new invading pathogens. This internal balance between the hosts

immune system and infection load is constantly changing in accordance to hosts health, external

16

changes and infectious processes. Healthy individuals are able to maintain a non-pathogenic

level of antigens. In immunocompromised individuals, however, the levels of already colonised

antigens can increase and lead to clinical symptoms (14, 15).

1.3 Synergistic and antagonistic impacts of

coinfection

In order to identify the factors that shifts an individual from healthy to immunocompromised, it

is necessary to look at the details of the interactions amongst pathogens and the hosts. When a

new pathogen is introduced to a host with an already established pathogen community, the

preestablished pathogen can use the host’s immune system to their advantage by either

propagating or suppressing the cytokine network and T lymphocyte subsets. This is dependent

on the pathogens interacting synergistically or antagonistically with each other (16, 17) (Figure

1.1). For antagonistic interactions, pathogen burden is generally supressed by an already

established antigen. In synergistic relations pathogen burden is enhanced, which tend to be the

most harmful for the hosts (8).

Figure 1.1 Over of potential consequences of multiple infections with regards to hosts manipulation

(2).

Synergistic interactions are seen in some of the most prevalent animal and human diseases (12).

Examples of synergistic interactions can be seen in HIV patients were the viral infection causes

immunosuppression in the host, resulting in susceptibility of secondary infections such as

17

tuberculosis. This leads to an increase of viral load through HIV-1 promotion (18). The

protozoan parasite Leishmania major is another etiologic agent which uses the host immune

response to its and other pathogens benefit. L. major takes over host translational machinery

through mTor signalling disruption, leading to higher parasitic infection susceptibility (19).

Synergistic co-infections tend to be more harmful to the hosts as individually each antigen is

usually non-pathogenic, but within a co-infection enhancement of pathogens occurs leading to

clinical severity. Antagonistic interactions may also occur, which tend to be due to ecological

reasons (13) or physiological reasons.

Infections which exhibit antagonistic interactions tend to be seen in helminth parasitic

infections. A well-known example is the phenomenon of concomitant immunity, seen in

schistosomes and Echinococcus granulosus (20). This form of antagonistic reaction is a result of

adult worms preventing establishment of larvae through eliciting an immune response within the

host. Schistosome adults evade this immune response by a specialised surface membrane which

obtain hosts proteins, whilst new larvae are eliminated by the immune response. (21, 22)

Another example is helminth co-infections with schistosomes or hookworm, commonly seen in

children which results in high levels of anaemia. This is due to ecological factors of helminths

which is the result from nutrition competition for blood and schistosome egg extravasation in

the bowel wall (12, 13). Timing of pathogen infection can cause different interactions,

pathogens which have colonised early will have a benefit over newly infecting pathogens. This

is due to the ability to increase or lower immune response. For example, if a pathogen has

already colonised at a target organ, this may induce antagonistic interactions between newly

arriving antigens which attempt to colonise at the same infection site (11).

Different parasitic species will affect the host’s immune system uniquely, in parasitic infections

were more than one species of parasite (polyparasitism hereafter) can be found, these

interactions can become quite complex. Protozoans tend to polarise responses towards the Th1

response (pro-inflammatory stimulus), whilst helminths induce Th2 (anti-inflammatory

stimulus) and regulatory T cells (17, 23-25). Recent studies also suggest that parasites establish

chronic infections by expanding regulatory T cells and polarising the immune system in favour

or counterbalance along the axis of Th1/Th2 or Th17/Treg in response to co-infecting parasites.

18

This ability to polarise the immune system in favour or against other parasites ultimately is

important to the host’s resistance, susceptivity and immunopathology (23).

1.4 Parasitic infections in wildlife

The existence of co-infection is beyond an individual animal, it can be seen to affect the wider

population as well. Wildlife populations are effective sentinels for the environment, their roles

in understanding natural parasitological patterns are vital for further research into parasitic

population dynamics. These dynamics have been a key focus in ecology for many years, though

much of the progress has been made in human infections and veterinary epidemiology rather

than in wildlife systems.

Whilst the effects of parasitic infections on ecology have been at the forefront of human,

livestock and veterinary research for many years, wildlife parasitology has not received the

same level of research which has resulted to a large gap of knowledge (26). The ecology and

evolutionary biology of wildlife disease is important for improvement of current disease

prevention mechanisms. This is predominant in ecological parasitology, which can expand on

parasitology effects population dynamics and evolutionary ecology literature. (27). Using

natural animal populations, such as reptiles, a vast amount of information on pathogen dynamics

can be discovered.

Reptiles are strong environmental sentinels due to their sedentary, terrestrial lifestyle and large

abundance in agriculture. Their omnivorous diets and long lifespans allow them to be exposed

to a variety of pathogens and chemicals allowing long term observations on pathogenic diseases

and cumulative toxins (28). The Australian Shinglebacks lizard Tiliqua rugosa, is an extremely

good example of an Australian environmental sentinel. Not only are they omnivores but they

also have strong interactions within rural and urban environments, allowing observational

differences.

Reptiles are known to be susceptible to a wide range of pathogenic agents, it is uncommon to find a

wild skink with one or no infection. Due to reptile's high susceptibility to infectious agents’ disease

is caused by a multitude of factors (6). Not many parasitological species have been well researched

in the Tiliqua rugosa population, though the few reports available are mainly focused on rickettsia,

ticks, blood parasites (6, 29) and ectoparasites Bothriocroton hydrosauri and Amblyomma limbatum

19

(30). Epidemiology tends to reflect the multifactorial nature of disease found in reptiles, resulting in

the large range of clinical outcomes. There are several factors interplaying, including environmental

conditions, diet, periods of activity and inactivity, refuge site availability, stress and external factors

like human interaction (31). Thus, creating a model can account for the numerous amounts of

interactions is impossible, though by looking at a few of these factors in accordance to disease

spread it allows a vast amount of information to be gained.

Even though the obvious difficulties in obtaining replicable data, reptiles prove to be good

indicators for changes in the environment, disease progression/patterns and the health of wide-

ranging ecosystems. Infections affecting reptiles have only recently been of importance to

researchers due to the ability to gain insight into Arboviruses such as yellow and dengue fever

which effect both reptile and human health (32). Further research into the Arboviruses in reptile

health has allowed further understanding of the epidemiology of such viruses.

1.5 Tiliqua rugosa

The focus of this study is the Australian Shinglebacks (Tiliqua rugosa) which acts as a good

environmental sentinel for Australian environment conditions. Tiliqua rugosa, commonly known

as the Shinglebacks lizard or Shingleback lizard are an Australian native reptile which inhabits drier

regions in Southern Australia all the way to coastal areas in Western Australia (33). It is a viviparous

large day-active skink which is known for its monogamous long-term partnerships and stable home

regions which overlap frequently with other Shinglebacks lizards (34). Shinglebacks lizards are

typically found to inhabit semiarid plains, with harsh summers and cooled winters (35). Shingle back

lizards are one of the largest skinks within Australia, with an average weight of around

600 to 900 grams and average snout-vent length of 16 to 18 inches. Shinglebacks have

distinguishable physical characteristics including their ‘pine-cone’ scales which encase its entire

upper body and its unique blue coloured tongue which is how the common name ‘blue-tongue’ lizard

originated from (36). The tail has a rough, ‘knobby’ appearance which is close in measurement to

the head of the reptile, this tail acts as fat storage which noticeably depletes as the Shinglebacks

loses body weight due to illness or other factors (34, 36).

Shinglebacks lizards are colonial in nature and have sedentary lifestyles, usually staying within the

same home ranges, which are approximately 200 meters in diameter (36). Studies looking into the

colonial behaviour of Shinglebacks has allowed great insight into Shinglebacks social-network

20

interactions and how they are affected by the presence of parasites, for example, ticks which benefit

on closer linked social-networks (29). These studies are great indications into how other parasites

and aerosol viruses can infect at such rapid rates due to shared refuge sites, feeding grounds and

home ranges.

Tightly bonded skinks benefit from shared refuge areas, through lowered predation and enhanced

thermoregulation by reduced surface area exposure. Individual Shinglebacks in comparison will

commonly be seen in separate refuge sites as transmission of parasites and aerosol illness are easily

transmissible when refuge sites are shared with more than one lizard. Research has found that an

abundance of directly transmitted parasites depends on how frequently direct contact is made among

hosts individuals, therefore, lizards will try to find refuge alone to lower chances of parasitic

transmission. The type of refuge area will be a contributing factor to parasitism in the population, in

higher foraging areas Shinglebacks will share refuge areas as a larger density of the lizard population

will be found in these areas. In comparison to drier arid areas were Shinglebacks have the ability to

refuge alone (35).

1.6 Parasitic infections in Tiliqua rugosa

Due to the multiple factorial nature of disease it is extremely difficult to understand the nature of

disease within a Shingleback’s community. There are several factors interplaying, including

environmental conditions, diet, periods of activity and inactivity, refuge site availability, stress and

external factors like human interaction. Thus, creating a model which can account for the numerous

amounts of interactions is impossible, though by looking at a few of these factors in accordance to

disease spread it allows a vast amount of information to be gained. The benefit of their long lives

allows for long term research into various fields of sciences, therefore expanding our knowledge of

disease pattern.

Parasitic infections in Tiliqua rugosa negatively impact on the individual hosts and the wider

Shinglebacks population. Such infections have proven to be more detrimental on adult males

than females (37), due to the adverse effects on overall host fitness. For male adults, lowered

hosts fitness is disadvantageous as it reduces home ranges, territorial behaviour and social

status. For Shingleback lizards, courtship and activity levels can be strong indicators of an

individual’s ability to cope with parasitic infections which directly relates to immunity,

resilience and better host fitness levels (38). Parasitic levels in a host has also shown to cause

21

issues with mating; female Shinglebacks will prefer to pair with unparasitised males, which is a

result of sexual selection were the female will choose males which may be able to transfer

strong immunity traits to their young (39). Hosts must make a trade-off between fitness and

defence against pathogens (40). Shinglebacks may sacrifice pathogen defence for increased

activity to maintain territory and expand home range for mating purposes. In doing so,

susceptibility to higher parasitic loads, parasitic diversity and opportunistic pathogens becomes

an issue for the Shingleback (38).

1.6.1 Protozoans

Under natural conditions, reptiles will harbour a variety of protozoan infections. Protozoans can

reproduce quickly, thus when a host is immuno-compromised protozoans are able to take

advantage and reproduce quickly without immunity interference. Due to this ability protozoans

can become clinically significant, especially in hosts which are immuno-compromised. These

volumes of protozoan parasites can cause host population dynamic changes (41). The most

commonly known protozoan in the Shinglebacks lizards are Hemolivia mariae, transferred

through ingesting ectoparasites (37). This blood parasite seems to cause no clinical issues,

although Shinglebacks with larger home ranges that cross over with others, tend to have higher

rates of infection (38).

Eimeria tiliquae is another protozoan from the Eimerridae family that infects Shinglebacks

lizards (43). This protozoan family was seen to infect 21% of the Shinglebacks entering

rehabilitation centres. Nycotherus trachysauri has also been seen in the rectums of T.

rugosa. A few other protozoan parasites have been discovered in the T. rugosa population

although speciation which is the formation of a new and distinct species was not completed by

the researcher (Table 1.1).

22

Table 1.1 Protozoans found in Tiliqua rugosa

Family Species Reference

Karyolsidae Hemolivia mariae Smallridge and Bull (2001a) (42)

Eimeriidae Eimeria tiliquae Yang., et al (2013) (43)

Nycotheridae Nycotherus trachysauri Johnston (1932) (44)

Lankesterllidae Schellackia sp O’Donoghue (1998) (45)

Trichomonadidae Trichomonas Johnston (1932) (44)

Balantiididae Balantidium spp. Yang., et al (2013) (43)

Bodonidae Bodo sp. Johnston (1932) (44)

Peranemataceae Copromonas sp. Johnston (1932) (44)

Entamoebidae Endamoeba sp. Johnston (1932) (44)

Table 1.2 Helminths found in Tiliqua rugosa

Cestode Oochoristica trachysauri MacCallum (1921) (46), Johnston

(1932) (44)

Nematodes Abbreviata Antarctica

Jones (1992) (47)

Oxyuris sp.

Thapar (1925) (48)

Thelandros trachysauri Johnston and Mawson (1947)

(49)

A.turnidocapitis Jones (1983) (50)

Phyalopteroides filicauda Jones (1985) (51)

Maxyachonia brygooi Mawson (1972) (52)

Thelandros trachysauri Johnston and Mawson (1947)

(49)

23

Parapharyngodon fitzroyi sp. Johnston and Mawson (1947)

(49)

Pharngodon tiliquae Baylis (1930) (53)

Kriseilla lesueurii Jones (1986) (54)

Pseudorictularia disparillis Irwin-Smith, (1922) (55)

Veversia tuberculata Johnston (1932) (44)

Trematodes

Brachylaima cribbi Butcher and Grove (2005) (56)

Microphallus sp. Angel and Mawson (1968) (57)

Paradistomum crucifer Angel and Mawson (1968) (57)

1.6.2 Helminths

Intestinal helminth parasites are extremely common in Shinglebacks lizards (24), the most

predominant issues caused by helminth infections is negative impact on home ranges. In

previous studies a high percentage of 89.4% of helminths were seen in the sample Shinglebacks

population, with an average of 1.3 eggs gm-1 of faecal mass. Males were more likely to carry

parasitic infection with a prevalence of 100% parasite infection score and the females had an

80.8% parasitic infection rate. T. trachysauri was the predominant (89.4%) parasite, but O.

trachysauri was also found, with (19%) male and (11.5%) females both presenting infections

Helminth infections seen in the Shinglebacks lizards are quite predominant (47), their presence

has been noted from the early mid 1900’s. Although, these studies did not focus on the types of

interactions the helminth may cause to the host and the population.

24

1.6.3 Viral infections

There are several important viruses infecting reptiles, although they do not stand out alone,

rather they play a part of the multi-factorial nature of disease seen in reptile hosts (32). There

are seven types of known viruses; the RNA viruses, specifically the Nidovirales, Coronaviridae

family is highlighted in this paper due to its similarities to the partially genomic characterised

novel Tiliqua rugosa Nidovirus (6).

1.6.3.1 Nidovirales

The Nidovirales possess the largest known RNA genomes (58); these viruses are all enveloped

non-segmented positive- sense RNA viruses (59). They are a large diverse group which

significantly impact both humans and animal health. Nidovirus pathogenesis ranges from mild

enteric infections to severe respiratory diseases, which include the severe acute respiratory

syndrome (SARS) (60). The Nidovirales contain unique families and subfamilies which vary in

genomic size. The Arteriviruses are associated as ‘small’ nidoviruses, whilst the Coronaviruses,

Toroviruses and Roniviruses are considered as ‘large’ nidoviruses (58). The Coronaviridae

family comprised of the Coronavirus and Torovirus genera have the largest identified RNA

genome containing approximately 30 kb genomes (61). This viral family has a unique ‘crown-

like’ spike protein capsid (100). The Coronaviridae virus has distinguishing genomic traits,

including multiple Open Reading Frames (ORFs) with a replicase gene with two overlapping

ORFs named 1a and 1b. When translated these ORFs produce polyprotein pp1a and pp1ab.

These key morphological features are also seen in the Nidovirale family which creates a

connection between the two Coronaviridae and the Nidovirales and allows physiological,

morphological and clinical pathogenesis to comparable between the two groups (62, 63)

The newly discovered Nidovirus effecting Shingleback lizards in Australia has presented similar

clinical and genomic traits to a previously researched Nidovirus in ball python snakes, which

allows good comparable researched data on a similar Nidovirus to this newly discovered Tiliqua

rugosa Nidovirus which has no previous data. The partial genomic characterisation of the

Tiliqua rugosa Nidovirus allows insight into the phylogenetic placement of the virus and its

25

genomic similarities to the Coronoviridae’s and the ball python and green tree snake Nidovirus .

(6, 61).

1.6.3.2 Coronoviridae

The Coronaviruses cause a large variety of significant diseases in animals and humans and its

relevance to future research is extremely important (61). SARS-CoV, a group of 2b

βcoronavirus are endemic in human populations causing approximately 15-30% of respiratory

tract infections in a year. In immunocompromised individuals, neonates and the elderly these

symptoms become severe causing upper respiratory tract infection (URTI) symptoms.

The Coronaviridae family has caused significant pathogenesis in the porcine, bovine and fowl

industry which causes substantial economic burden. The cattle industry has Bovine CoV and

infectious bronchitis virus (IBV) which causes mild to severe respiratory tract infection in cattle.

This virus is significant due to its ability to easily cross over to other ruminants, causing

diarrhoea, weight loss, dehydration, lowered milk production and depressive behaviour (64).

IBV also takes its toll in the fowl industry by significantly diminishing egg production and

weight gain, this has caused substantial losses in the fowl industry each year (61, 64). The

porcine industry has the porcine reproductive and respiratory syndrome virus (PRRSV) which is

the costliest viral pathogen affecting the pork industry. Its ability to cause clinically significant

disease, maintain long lived infections and evade innate and adaptive immune responses make it

extremely difficult to control and eliminate (65). Nidovirales cause reproductive failure, poor

growth performance and respiratory illness and fatality in young piglets (66). In adult pigs, this

virus presents as a subclinical disease which participates as a co-factor in various polymicrobial

disease syndromes (65).

26

Figure 1.2 Maximum likelihood phylogenetic tree of Nidovirus Tiliqua rugosa (1)

27

1.6.3.3 Pathogenesis of the novel Nidovirus in Tiliqua rugosa

The ‘Shinglebacks’ Nidovirus has been noted since 1990 and has caused an influx of sick

Shinglebacks to enter rehabilitation centres, showing clinical signs of upper respiratory tract

infections (URTI) (6). The respiratory disease in ball pythons shares many similarities to the

Shinglebacks flu; including its close genomic relations it also shares very similar characteristics

in terms of clinical symptoms (67). Similar strains of the Nidovirus in the ball python and green

tree python allow some insight into the genomics, virology and pathogenesis of the novel

Tiliqua rugosa Nidovirus (60, 68).

The python Nidovirus causes morbidity and mortality in both wild and captive reptiles. The

pathogenesis seen in these snakes all presented pneumonia characteristics, including the

enhancement of mucous production in the airways, foveolar spaces, trachea, internal choanae

and causal air sacs (101) These mucous characteristics are common clinical signs seen in the

Shinglebacks lizards, which could be an indication of similar pathogenic processes occurring (6,

68).

Clinically there is a significant reddening of the oral pathways in the Shinglebacks, which was

also seen in the Ball Python post mortem findings. The redness seen in the Shingleback lizards

usually lead to discolouration of the gum, where in severe infections the colour would be a

white or grey colour (6, 67). The progression of this virus in an untreated host leads to open

mouth breathing due to an increase in respiratory effort. This ‘gasping for air’ is due to the poor

gas exchange occurring from morphological changes causing thickening of the lungs air ways

and alveoli in infected Shinglebacks. Finally, the host will lose significant weight due to

inappetence. In Shinglebacks, this is clear when the tail becomes flat or deflated and the lizard

loses its aggressive behaviour, ultimately this will lead to anorexia and death, which has also

been observed in Ball python Nidovirus symptoms (6, 67).

Research is still progressing into analysing this new clade of Nidovirus which are affecting

reptiles (69, 70). These studies progress into understanding the relationship of the virus to

pneumonia and if the high mortality rates are due to a causal relationship or if it is due to a

complex, multifactorial natured disease. Some other factors that may play into the multifactorial

28

nature of reptile disease are husbandry, age, immune status, sex and co-infections or

opportunistic pathogens (6, 67)

Figure 1. 3 . a= healthy Shinglebacks body condition, b= mild URTI body

condition, c= severe URTI body condition. Picture provided by Tasha

Henning’s , hospital manager, Kanyana Wildlife Rehabilitation n Centre.

a

b

c

29

1.7 Aims and Hypothesis

The main aims and hypothesis of this study is to investigate if any association exists between the

abundance and diversity of individual parasitic loads, with the presence/absence of the novel

Tiliqua rugosa Nidovirus.

Objective

1. To obtain baseline data on the prevalence of viral infection and parasitic infection

2. Confirm the proportion of animals infected by viral, parasitic and co-infections.

3. Examine the possible interactions seen in co-infected Shinglebacks.

Hypotheses

1. Shinglebacks will have a high prevalence of viral and parasitic infection.

2. The abundance and infra-community diversity of gastro-intestinal parasites will

be greater in shingleback (Tiliqua rugosa) lizards which are co-infected with the

Shingleback Nidovirus

3. Co-infections will have a strong positive correlation to Shinglebacks which test positive

for Viral symptoms

30

Chapter 2 Methodology

2.1 Sample collection

All samples were collected through the Native Animal Rehabilitation Centre (Native ARC) and

Kanyana Wildlife Rehabilitation Centre. Permission of the board of each centre was given

before sampling began and with the approval of the animal ethics committee from Murdoch

University (20180205). All samples were taken from shingleback lizards from April 2018 to

April 2019, with each shingleback lizard being placed into a ‘sick’ or ‘healthy’ category based

on veterinary assessment on the shingleback’s clinical symptoms. Each Rehabilitation centre

followed their own health assessment protocols for the evaluation of the shingleback flu.

Shinglebacks lizards were assigned categories based on presentable clinical signs such as, large

mucous build up in oral cavity regions, mucopurulent oculonasal discharge and reduced body

condition (6). ‘Healthy’ shinglebacks’ did not present signs of upper respiratory tract infection

(URTI), though usually had other factors such as injuries received from cars or pet incidences.

All shinglebacks’ which past the assessment for euthanasia (per Kanyana and Native ARC

protocol) was treated accordingly to their symptoms on admission (71).

All samples were taken with supervision of a veterinarian (Native ARC) or trained handlers

(Kanyana), with minimal animal stress procedures for data collection. All animal handling

procedures required for mouth swap samples were in accordance with Parks and Wildlife

standard operating procedures

Case definition was established to assign individuals to categories of ‘symptomatic’ and

‘asymptomatic’. Symptomatic Shinglebacks demonstrated signs of oculonasal discharge (Figure

1.1). Asymptomatic shinglebacks where considered healthy and had no signs of URTI (this

included any shinglebacks with trauma from injuries). Samples were collected at point of

admission (or as close to admission date). Strict asepsis techniques were adhered to in the

31

handling and collecting of samples, to eliminate potential cross-contamination. With the aid of

Shingleback handlers, the lizards were restrained and a sterile (15 cm) single-ended cotton tip

applicator was used to collect secretions from the glottis area of the oral cavity. The cotton tip

was placed into 0.5 ml of Viral Transport Media (Medium 199 + Penstrep + Fungizone; Sigma)

(6). The sample was immediately placed into an esky at approximately -20°C and then

transferred into a -80°C freezer within a two-week period at the Murdoch Parasitology

Laboratories.

Biological data was collected on each lizard as per each rehabilitation protocol and included

age, weight, snout-vent-length, tail length, eye/oral discharge, colouration, level of aggression

towards handlers and general body condition. All animals were examined by a veterinarian (as

per rehabilitation protocol) and were monitored thereafter by experienced staff for clinical signs.

The principal investigator (PI) did not take their own measurements but used both

Rehabilitation information for later analysis.

Faecal samples were taken when naturally produced and analysed on the same day at Murdoch

Parasitology laboratories. This was usually within the in the first four days of the Shinglebacks’

admission into the Rehabilitation Centre, prior to the administration of anti- flagellate treatment

as per Kanyana protocol. Samples were taken post treatment if the animal did not naturally

defecate prior.

2.2 Measurement of age and body mass

The Shingleback’s entered into this study were categorised by several factors, one of

which being age and weight. Age was calculated by a body condition index which

compared weight of the Shinglebacks in accordance to the snout-vent-length (Table 2.1).

Shingleback categorised as underweight or obese were given treatment in accordance to

aid in achieving a healthy weight. The calculation of age using length and weight was

directly taken by Kanyana Standard Operating Procedure.

32

Table 2.1. Age and body mass estimation per the weight and length of the Shinglebacks.

Information access provided by Tasha Hennings, hospital manager, Kanyana Wildlife

Length of Shinglebacks (Snout to end of tail)

< 20cm- 26cm 26cm- 32cm 32cm- 40cm +

400 – 600g + x Fat/ Obese Adult (4 years +)

2.3 Measurement of body condition

Body condition was measured through the residuals of snout-vent-length and weight linear

regression, performed through R studios, ggplot package (72). The body condition gives an

indication of possible effects of viral infection on the body mass of the Shinglebacks.

2.4 Measurement of viral symptom score

The measurement of viral symptom score was performed by a score of 1 and 0 (1=signs of

discharge, 0=no signs of discharge), which was based on observable mucopurulent oculonasal

discharge. These scores were based on vet’s diagnosis of the flu (Figure 2.1).

33

2.5 Faecal Analysis

Parasitology examination to detect gastro-intestinal (GI) parasites was processed through faecal

samples via traditional zinc sulphate faecal flotation and direct smear parasitological techniques

(5). Faecal samples were collected as soon as produced, to analyse when still fresh. All samples

were weighed, with 2g being the maximum weight used. Some samples were too small and

thus; samples with less than 2g of weight were calculated accordingly to its weight. Faecal

samples were then analysed by a direct smear for motile parasitic entities and with quantitative

zinc sulphate sedimentation flotation technique as per, with small alterations as seen below.

2.5.1 Faecal Smear

Faecal smear was used for the identification of mobile protozoans (Trichomonads, amoeba)

which float poorly or can become distorted by floatation solutions. Faecal mass is mixed with

saline solution and homogenized. Small droplets are placed onto the microscopic slide with 2- 3

drops of malachite stain (4). For Microscopic analysis coverslip is placed on top of solution

mix. Microscopic examination will require use of 10x to 40x magnification (5).

Figure 2 .1. a= eye discharge, b= mucous in oral cavity. Picture provided by Tasha

Henning, Kanyana Wildlife Rehabilitation Centre.

a

b

b

34

2.5.2 Faecal Flotation

Faecal flotation is used commonly in veterinary and research practices, this procedure

concentrates parasitic eggs and cysts and removes debris. This method is based on the principle

that density in parasitic entities is less than that of the flotation medium.

Using an accurate weight scale, weigh approximately 2g of faecal sample. The faecal mass must

be homogenized with Zinc Sulphate (gravity of 1.2) till it reaches 10 ml in volume (73).

Centrifuge for 2 minutes at 2000 rpm, using a centrifuge balance if needed to level out the

centrifuge. During the centrifuge time frame a Bunsen burner will be required for the

sterilisation procedure on an inoculation loop (74, 75). Following (76) procedure retrieve

approximately 2-5 lobs of suspended inoculum and place them onto a 76.2 x 25.4 mm

microscopic slide. Place a 22 x 22 mm coverslip on top of the solution and analyse the sample

(5).

2.5.3 Analysis

All faecal samples were screened using an Olympus BX50 microscope and images taken using

the CellSens Standard micro imaging software. Parasite species were identified using

magnifications of x10 to x100. Analysis of all samples was undertaken by the PI avoiding the

risk discrepancy between multiple technicians.

2.6 Virology

For detection of shingleback Nidovirus 1 RNA in swab samples, the methodology of (6) was

followed. Briefly, RRT-PCR was performed using the primers XYZ and the probe ABC.

Primers were used at a concentration of 900nM and probe at 450nM using AgPath- ID One-Step

reaction mix (Ambion). Reaction conditions were as follows, 45°C for 10 min, 95°C for 10 min,

followed by 45 cycles of 95°C for 15 s and 60°C for 45 s.

35

2.7 Statistical analysis

Parasite and viral prevalence’s were expressed as a percentage of infected hosts, with

confidence intervals calculated assuming a binomial distribution. Confidence intervals around

mean intensities of infection (number of parasites per infected host) were calculated by

bootstrapping. Associations between categorical variables were examined with Chi-squared or

Fisher’s exact test. Differences in the means of continuous variables were tested using analyses

of variance or, if the assumptions of normality of residuals and homogeneity of variances were

not satisfied, by a non-parametric Kruskal-Wallis test. All statistical analyses were performed

using R (72) or QPweb (77).

36

Chapter 3 Results

3.1 Shingleback demographics

Ninety-three Shinglebacks were tested in this study for Nidovirus and parasitic infection. Of the

93 lizards sampled, 57% (n=53) were juveniles and 43% (n=40) were adults. Shinglebacks were

found in a variety of locations (Figure 4.1) and entered rehabilitation centres for signs of flu, car

accident, lawn mower injury or animal attack.

Figure 3.1 . Distribution of the original locations in which Shinglebacks lizards were found before en tering rehabilitation centres. Dot size indicates number of animals collected at a location.

37

Head length of sampled Shinglebacks varied from 1.9 cm to 9.8 cm, and body weight from 33 g

to 773 g, with a strongly positive relationship between length and weight (Figure 3.2).

Figure 3.2 Relationship between snout-vent-length and weight in the sampled Shinglebacks.

Regression line: Weight = -130.8 + 86.2SVL, R2 = 0.49, F1,91 = 88.0, P < 0.0001.

3.2 Parasitology

Faecal analysis identified three protozoan genera and three nematode genera in Shinglebacks sampled, with

a number of other protozoan cysts and nematode eggs that could not be

Faecal analysis identified three protozoan genera and three nematode genera, with a number of other

protozoan cysts and nematode eggs that could not be identified further (Figure 3.3; Table 3.1). The

percentage of Shinglebacks infected with any GI parasite was 73.1% (n=68), with 57% (n=53) infected

with protozoans, 49% infected with helminths and 35.5% (n=33) infected by both parasite groups identified

further (Figure 3.3; Table 3.1).

Figure 3.3 a - h. Parasitic species found during faecal analysis: a.1= Sporulated Eimeria SP.; width: 7 μm , length: 6 μm , magnification:

40 x, a.2= Unsporulated Eimeria sp.; width: 5 μm , length, 5 μm , magnification: 40x, b= Entamoeba sp.; width: 6 μm , length: 10 μm ,

magnification: 40xc= Trichomonas sp.; width: 5 μm , length: 7.2 μm , magnification: 40x, d = Unknown protozoan cysts; 7 μm, length 6

μm, magnification: 40x , e= Oxyurid sp.; width: 18 μm, length: 12 μm, magnification: 20x ; f= Physaloptera sp .; width: 15 μm , length: 17

μm , magnification: 40x, g= Unknown nematode eggs; width: 23 μm, length:12.1 μm, magnification: 20x.

40

39

Table 3.1 Prevalence (%) and mean intensity of parasite taxa found in sampled Shinglebacks

Protozoa

Parasite

Phylum

Parasite

Family

Parasite

Genus or group

Prevalence (%)-

(95% confidence

interval)

Mean intensity -

(95% confidence

interval)

Apicomplexa Eimerridae Eimeria sp. 40.9 (31.1-51.1) 9780 (4730-18600)

Amoebozoa Entamoebidae Entamoeba spp. 8.6 (4-16) 43.2 (16.2-113)

Parabasalia Trichomonadidae

Trichomonas

spp.

26.9 (18.7-37.0) 1520 (798-2780)

Helminth

Unknown

Unknown

Unknown

protozoan cysts*

35.5 (26.3-45.7)

1260 (734-2010)

Nematoda Oxyuridae Oxyurid spp. 44.1 (34.4-54.3) 314 (135-794)

Nematoda Physalopteridae Physaloptera sp. 3.2 (0.9-9) 440 **

Nematoda Spiruroidae Spirurida spp. 1.1 (0.1-5.7) 1**

Nematoda Unknown Unknown nematode

eggs*

3.2 (0.9-9) 21.5 (3-21.5)

*Unable to classify parasitic species further

** Only one sample analysed, therefore no confident intervals

40

There were no significance differences in prevalence between adult and juvenile lizards for any

of the parasitic genera (Figure 3.4: for Eimeria, , χ21

= 0.02, P = 0.88, for Entamoeba, FE test, p

= 0.46, for Trichomonas, χ21= 0.35, p = 0.56, for protozoan cysts χ2

1 = 0.27, p = 0.6, for Oxyurid

, χ21

= 2.012, p = 0.15, for Physaloptera, FE test, p = 0.26, for Spirurida, FE test, p = 1, for

nematode eggs, FE test, p = 1).Similarly, for those parasites for which intensities of infection

could be determined, there were no significant differences between adult and juvenile lizards in

mean intensity (Table 3.2: Kruskal-Wallis test was used for analysis: for Eimeria sp, χ21= 0.06,

p = 0.81, for Entamoeba, χ21= 1.21, p = 0.27, for Trichomonas, , χ2

1 = 0.04, p = 0.84, for

protozoan cysts , χ21

= 0.24, p = 0.62, for Oxyurid, , χ21= 0.35, p = 0.56 ).

Figure 3 . 4 Prevalence of p arasit e taxa in adult and juvenile Shinglebacks . Black lines are 95 % confidence

inter vals.

41

Table 3.2 Mean intensity parasite taxa in adult and juvenile Shinglebacks (95%) confidence interval in

parentheses

Figure 3.5 Mean number of parasites/individual host in adult and juvenile

Shinglebacks.

Eimeria sp. Entamoeba

sp. Trichomonas sp. Protozoan Cysts Oxyurid spp.

Juvenile

12400 52.7 1770 1460 257

(5550-24500)

(17-140) (744- 3920) (736-2580) (85-553)

Adult

6240 (1170-

15 1210 966 386

21300) (10-15) (373-2570) (263-2380) (81.7-1500)

42

The mean number of parasite taxa/individual host over all sampled Shinglebacks was 1.6 (95% confidence

interval 1.3-1.9), with no difference between adult and juvenile lizards (Figure 3,5; Kruskal-Wallis test, z

= 0.07, P = 0.94).

3.3 Virology

Nidovirus infections were found at a prevalence of 59.3% (95% confidence interval of 48.969.3%).

There was no significant difference between prevalence in adult and juvenile

Shinglebacks (Figure 3.6, χ21 = 0.85, p = 0.36).

Figure 3.6 Prevalence of Nidovirus l infection in adult and juvenile Shinglebacks. Black lines indicate 95%

confidence intervals.

43

3.4 Co-infection

Co-infections with parasites and Nidovirus were common within the sampled Shinglebacks

population, with 48.4% of animals carrying both parasitic and viral infections, 26% carrying

parasitic infections only, 9% carrying viral infections only and 17% with no infections of either

parasites or Nidovirus (Figure 3.7). The prevalence of co-infections did not significantly alter

between adult and juvenile Shinglebacks (table 3.3, Kruskal-Wallis test, χ21

= 1, p = 0.32).

9 %

26 %

48 %

17 %

Viral infection Parasitic infection Viral and Parasitic infection

Figur e 3 . 7 Prevalence (%) of viral and parasitic infection s in

Shinglebacks

Figure 3.8 . Percentage Prevalence of Viral Infection in Accordance to Parasitic Infection

P value < 0.5

44

Percentage prevalence of viral infection was significant in comparison to the presence or absence

of parasitic infection. There is clear correlation between co-infection resulting in high percentage

prevalence of clinical symptoms (figure 3.8, χ21= 9.3, p = 0.02).

Number of parasitic species seen was statistically significant in comparison to viral prevalence (Figure

3.9). Viral presence had a strong correlation with parasitised hosts which harboured parasitic infections

ranging from 1 parasite to 4 parasites. (Figure 3.9, χ21

= 5.15, p = 0.02).

Figure 3. 9 Mean number of parasite t axa in Shinglebacks with and without Nidovirus infections

45

3.5 Clinical signs of viral infection

There was a strong association between viral infection and the presence of discharge from the eyes

or mouth (Figure 3.10; χ21 = 11.1, p = 0.001).

Body condition score, however, did not differ between Shinglebacks with and without viral infection

(Figure 3.11; F1, 89 = 0.7, p = 0.41).

Figure 3 .10 Percentage prevalence of viral infection in accordance to viral symptom score

P <0.5

Clinical signs

46

Figure 3.11 Mean body condition score of Shinglebacks with and without Nidovirus infection.

There was a significant correlation seen between viral symptom score and body condition.

(Figure 3.11, χ21= 4.6, p = 0.033). The differences between groups can be seen by the overall

score of Shinglebacks with higher body conditions in the population with no clinical URTI

symptoms.

For those Shinglebacks which had a Nidovirus infection, clinical signs of infection (discharge

from the eyes or mouth) were more likely to be present if animals were also coinfected with at

least one parasite species (Figure 3.12; χ 21 = 4.5, p = 0.03); this relationship between parasite

infection and discharge was not present in animals that were not infected with Nidovirus (Figure

3.12; χ 21 = 2.7, p = 0.1). Furthermore, for those Shinglebacks infected with Nidovirus, the mean

number of parasite taxa was significantly greater when clinical signs of infection were present

(Figure 3.13; FE1,52 = 5.4, p = 0.02)

47

Figure 3.12. Percentage Prevalence of Viral Symptoms in Accordance to Parasitic Infection

P value < 0 .5

Clinical signs present Clinical signs present

Figure 3 . 13 Mean number of parasite taxa for virally infected Shinglebacks that

showed or didn’t show clinical signs

48

Chapter 4 General discussion

This is the first comprehensive study of gastro-intestinal parasitism in wild populations of

Tiliqua rugosa and the first study of co-infections of gastro-intestinal parasites and Nidovirus.

There were three main findings from this study. First, prevalence of infection was relatively high,

with 73 % of Shinglebacks infected with at least one species of gastro-intestinal parasite and

58.1% of Shinglebacks infected with the T. rugosa Nidovirus. Secondly, co-infections with

gastro-intestinal parasites and Nidovirus were common; 48.4% of Shinglebacks carried both

parasitic and viral infections, while 26% had only parasitic infections and 9% had only viral

infections. Finally, I found evidence that the major clinical signs of viral infection, discharge

from the eyes or nose, were enhanced in Shinglebacks which were coinfected with the virus and

gastro-intestinal parasites.

4.1 Prevalence of parasitic and viral infections

Parasitic infections are ubiquitous in wild animal populations (12) usually without the expression

of clinical disease (24). Compared to other vertebrate host groups, the parasitic fauna of reptiles

has been poorly studied (78), but nevertheless it is common for reptiles to carry a variety of

parasitic infections, particularly protozoa and nematodes, and these often appear to be non-

pathogenic (79).

A study examining gut content of Scincidae in Australia found Gastro-Intestinal (G)I parasites at

an infection rate of 16.9% out of the total population tested. In this sampled population the

Egernia group displayed higher infection rate of 80%, speculated to be due to its dietary

lifestyle (80). Omnivorous reptile species tend to carry heavier infections due to their diet. This

style of dietary lifestyle is also seen in the Shingleback lizards’ diets (80).

In the current study, oxyurids (pinworms) were the most common form of helminth infections,

with a prevalence of 44.1%. Oxyurids are frequently the most common endoparasite found in

reptiles , especially in lizards and turtles (81) although limited information is available on

49

pinworm infections in Shinglebacks. Due to their direct life cycle, high numbers of oxyurid eggs

are characteristically seen in reptile faecal mass. Oxyurids tend to develop a commensal

relationship with the hosts, with little pathology, although compromised immune systems due to

other infections or changed environmental conditions can lead to increased numbers and cause

obstructions or lesions in the gastrointestinal tract (47). Currently Abbreviata antarctica and

Thelandros trachysauri are the only identified nematodes under the Oxyurid family in the

Tiliqua rugosa. Hugh I Jones (1992) study on the Shingleback lizards discovered an infection

prevalence rate of 35% for A. anatarctica and 60.9% in T. trachysauri.

Other helminths identified in my study were the nematodes Physaloptera sp. and Spirurida sp. Eggs

of Physaloptera sp. have previously been reported in Shinglebacks (82), although little is known

about the effects of this helminth infection in reptiles. Parasites of the Physalopteridae family occur

ubiquitously in Australian reptiles, including skinks (83) with large densities of infection and often

co-infections with other parasites (83).

I did not find any evidence of infection with the cestode Oochoristica trachysauri, although it

has previously been found in Shinglebacks in South Australian region at a prevalence of 15%

(84). Oochoristica may not have been found in this study sample population due to the need for

an intermediate host to complete its life cycle. Due to beetles being required for Oochoristica to

finish its life cycle it would be less visible in faecal samples and may be a result to the lack of

Oochoristica in this study (84).

Protozoan parasites were extremely common in the sampled Shinglebacks population, with most

individuals testing positive for at least one protozoan genus. The most common protozoan

parasite was a species of Eimeria, at a prevalence of 41%. Yang et al. (2013) previously

described Eimeria tiliquae in Shinglebacks, at a prevalence of 21%.

Nothing is known of the life cycle or pathogenic effects of this species. Although eimeriids infect

a large range of hosts, often causing severe clinical disease called coccidiosis (85), little is

known regarding this family of parasites in reptiles. In Australia, the only other described

eimeriids in reptiles are three species of Isospora in Australian Lacertilia(43).

50

Other protozoan parasites found in the current study were Trichomonas sp. and Entamoeba spp.,

which Yang et al, (2013) discovered within the Tiliqua rugosa population at a prevalence of 35%,

similar to the prevalence 26.9% rate seen for the present study. There is a paucity of information

on trichomonas infections in reptilian hosts, although they are common in birds and mammals,

where they may be pathogenic in the urogenital, GI or upper respiratory tracts (86).

Entamoeba spp. was found in Shinglebacks in this study, at a prevalence of 8.6%. Entamoeba

spp are commonly seen in reptiles (87). The most common species is E. invadens, which causes

gastroenteritis in lizards and snakes, with clinical signs including anorexia, weight loss, vomiting,

mucoidal or haemorrhagic diarrhoea, and death (88). Other species of Entamoeba found in

reptiles are E. terrapine, E. insolita and E. ranurum (87). Entamoeba moshkovskii is found in

anoxic sediments and brackish coastal pools in Australia and may infect mammals, reptiles and

humans (89).

Nidovirus was present in 59.3% of Shinglebacks sampled in the current study, although this may

be an underestimate of the true prevalence as shedding of the virus may have stopped before

oral swabs were taken. Nidovirale infections seem to be long lasting with prolonged shedding

(67), although this has yet to be shown to be the case for the virus infecting

Shinglebacks.

Although the Nidovirus found in Shinglebacks is genetically distinct, it appears to be closely

related to the ball python Nidovirus (1). The ball python Nidovirus causes morbidity and

mortality in both wild and captive reptiles. Clinical signs in reptiles infected with the ball python

Nidovirus include the loss of type I pneumonocytes in capillary walls and the replacement of

type II pneumonocytes, which enhances mucous production (68). Increased mucous production

is also a common clinical sign of infection in Shinglebacks, which could be an indication of

similar pathogenic processes occurring (1, 68). Clinically, there is a significant reddening of the

oral pathways in Shinglebacks, which is also seen in the post mortem findings from ball python

Nidovirus infections; this is due to ventral oral swelling, caused by small mucosal haemorrhages

(67). In reptiles infected with the ball python Nidovirus, hyperplastic changes in the trachea,

oesophagus and oral cavities, with associated mononuclear and granulocytic interstitial

51

inflammation and epithelial necrosis (9), causing lung thickening, pulmonary haemorrhages and

upper and lower lesions in the respiratory tracts (60). The progression of this virus in an

untreated host leads to open mouth breathing due to an increase in respiratory efforts (60). This

‘exhaustion for air’ is due to the poor gas exchange occurring from morphological changes,

causing thickening of the lungs air ways and the alveoli, which is also present in infected

Shinglebacks (67). Finally, the host will lose significant weight due to inappetence (67). In

Shinglebacks, this is clear when the tail becomes flat or deflated and the lizard loses its

aggressive behaviour, ultimately this will lead to anorexia and death, which has also been

observed in Ball python Nidovirus infections (6). This loss of weight has been seen in

Shinglebacks demonstrating flu like symptoms, as seen in Figure 1.3, Shinglebacks which are at

the later phase of the viral infection will be nearly non responsive, have no visible fat left and

have grey pale gums with discharge in oral and nasal passages.

4.2 Co-infections with parasites and Nidovirus

Co-infection between gastro-intestinal parasites was common in this study, with almost half the

population carrying both infections. Co-infection prevalence stayed relatively the same within the

adult population in comparison to the juveniles.

Interactions between the GI parasites seen and the Nidovirus seem to be synergistic in nature,

which may be a result of immune modulation by the virus or parasite, resulting in enhancement

of the other. Viruses tend to have an immune-depressive effect which results to the down

regulation of cytokines. These cytokines are required for the immunity against protozoans (17).

An example of the immunosuppressive effects of helminth and viral infections can be seen in the

lymphotropic virus type 1 (HTVL-1) which enhances Stronglyoides stercoralis infections in the

hosts (90). Other evidence of helminths increasing the severity of viral infection is seen in

Hepatitis B, were virally infected hosts can obtain liver damage due to Schistosoma mansoni

parasitic infections. Schistosomes have been noted alongside Hepatitis B in other studies,

causing the enhancement of viral infections (17).

52

Another example of synergistic interaction seen between parasitic and viral infections is in

coinfections between leishmania and human immunodeficiency viruses (HIV) (91). The

exacerbating effects of HIV is due its ability to immunomodulate the hosts. HIV results in a

decreased rate of CD4 T cell count whilst promoting Th2 response. Macrophage effector and

antigen- presenting function were impaired and a decrease in reactive oxygen species production

and pro-motes IL-10 secretion (92). These immune modulations favour parasitic persistence,

therefore in patients with this co-infection present higher viral loads and faster HIV progression

of infection due to the immunological stress of the synergistic interactions between these

pathogens (93).

This study cannot confirm whether the cause of the respiratory disease is due to the infection of

the virus alone or the combination of infections with parasitic entities. Nonetheless, it is no ticeable

that a pattern emerges from this study, with the vast amount of population carrying both

infections. Though the percentage of co-infection is significant other factors such as stress, other

pathogens and environmental factors may come into play when formulating the possible reasons

for the severity of the Nidovirus infection.

4.3 Co-infections and viral clinical signs

Looking at the interactions between the viral infection with parasitic entities it is predominant that

the reason behind the death rates is due to the inability of the hosts to maintain such high

immunological demands. This could be due to the synergistic interactions causing immune

manipulation to benefit the parasitic entities, or due to the inability to recovery from the initial

immunosuppression caused by the virus.

Parasitic infections within themselves have unique interactive patterns. Protozoan and helminth infections

can sometimes cause amplification of pathogenesis, especially in the case of poorly maintained pet reptiles.

Protozoans and helminths target different areas of the host’s body and activate different subsets of the

immune system. The imbalance caused by both parasitic groups trying to manipulate the immune system

can it self cause pathogenesis as the host is unable to stabilise a balanced immune system, this leads to

advantageous conditions for pathogen reproduction and colonization.

53

One of the most commonly studied parasitic interaction involving both protozoan and helminth

infections are between trypanosomes and Trichinella spiralis (94). These parasitic infections

tend to cause immunosuppression allowing the other infection to enhance within the hosts,

although this progression of infection can be sensitive to the time of infection (94). In a similar

study using research mice an increase of infection was seen in co infections between

Schistosoma mansoni and Toxoplasma gondii (95). Further clinical analysis of this co-infection

revealed sever liver damage, weight loss and higher mortality rates after several weeks with the

coinfection (95).

Interestingly, there was no relationship between the body condition of Shinglebacks and either

infection with GI parasites, infection with Nidovirus or clinical signs of viral infection. This may

be attributed to two factors, firstly snout-vent length was unable to be calculated for all

Shinglebacks. Therefore, head length did not act as a good measurement indicator for the effects

of the flu on Shingleback body weight. Secondly nearly all Shinglebacks in this study were of

poor body condition (healthy bobtails may have had car injuries, dog bites or other non-flu

sickness). Further research which includes wild populations with bigger dataset will allow for a

better understanding on the effects of the flu on body condition.

As hypothesised at the outset of this study, I found evidence that co-infection with GI parasites enhanced

the clinical signs of viral infection. Shinglebacks infected with both Nidovirus and GI parasites were

more likely to show oral or nasal discharge than Shinglebacks infected with

Nidovirus, without GI parasites.

The observable clinical signs associated to the Nidovirus is excessive mucous found in the oral

cavity, sneezing, serous to mucopurulent discharge from the eyes and nose, lethargy due to

inappetence, greying/whitening of the mucous membrane, depressive behaviour and the loss of

body condition, especially the tail area (1). These symptoms may be a result of possible synergistic

interactions, caused by the combination of GI parasites and the Tiliqua rugosa Nidovirus. Due to

this study being the first baseline study on the associations on internal parasites with the Nidovirus

in Shinglebacks, no literature is available for comparison to this study. Using previous literature

on viral infection and parasitic interaction, similar patterns of pathogen enhancement can be seen.

54

In previous studies with experimental mice, Trypanosoma cruzi infections were more severe in

mice with a viral co-infection. Another example can be seen in the murine leukaemia virus

(MuLV), which results in the enhancement of T. cruzi infections (96). This is due to T cells

being unable to respond to the T. cruzi infection, suggesting immunodepression on part of the

virus (96). Many more cases have been noted were viral infections are enhanced by protozoans

due to immunosuppression being caused by one of the entities (17).

One of those cases can be seen in honeybee colonies effected by Varroa destructor and pathogenic

deformed wing virus (DWV), which leads to the collapse of the colony due to host

immunosuppression (97). This form of suppression is seen by the marked transcriptional

downregulation of a member of the NF-kB gene family. This indicates that pathogen-parasite

interactions can restrict a variety of immune responses regulated by this transcription factor.

Therefore, this interference allows antiviral defence to be manipulation by pathogen interactions

(98). This suppression effect on the immune system is driven by viral replication within the hosts,

since Varroa does not seem to cause immune suppression alone (97).

4.4 Study limitations and further research

In summary this paper gives indication that within the Tiliqua rugosa population the Novel

Nidovirus plays a part of the multi dynamic nature of disease found in reptiles. Co-infection

with gastro-intestinal parasites seems to cause a synergistic effect alongside the virus, resulting

into observable clinical symptoms. Though further research in immunology and pathology needs

to be continued, to assess the immunological interactions occurring and asses the upper

respiratory tract for any potential signs of parasitic obstruction. Furthermore, parasitology work

must continue as blood parasites and ectoparasites were not examined for in this paper, which

may also be causing immune imbalance allowing for the progression of clinical symptoms.

From assessing information from Mark O’Dea et al, 2016 (6) the Nidovirus in the Shinglebacks

is perhaps acting alongside another pathogen or in combination with a multitude of comorbidities

in an individual to results into the upper respiratory syndrome seen in the

Shinglebacks.

55

Stress is a prominent factor not catered for in this study, but it plays a huge underling issue in

terms of factors that may progress clinical severity. All Shinglebacks entering the rehabilitation

centre have either serious injuries or been handled by humans. These stresses, which include

being caged for days or weeks can cause extra stress to the Shinglebacks, making clinical severity

more prominent when inspection were undertaken by the veterinarian diagnostic team.

Reptiles also have a heavy bacterial system, which could be another pathogen yet to be explored

(99). As well, no post mortem studies have been completed for Shinglebacks lizards, which will

allow a clearer understanding of the adult parasites which inhibit the host and see if any

obstructions or deteriorations of internal organs took place which could have led to the death.

56

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61

Chapter 6 Appendix

62

Figure 6.1. White gum commonly seen in Shinglebacks infected with URTI infection. Picture

provided by Tasha Hennings, hospital ma nager, Kanyana Wildlife Rehabilitation Centre.

a

Figure 6. 2 . a= Thickened discharge in eye. Picture provided by Tasha Hennings, hospital

manager, Kanyana Wildlife Rehabilitation Centre.