Pseudoloma Neurophilia: Progression of Infection and Transmission Characteristics of a Microsporidian Parasite in a Model Vertebrate, Danio Rerio

AN ABSTRACT OF THE DISSERTATION OF

Justin L. Sanders for the degree of Doctor of Philosophy in Microbiology presented on September 3, 2013. Title: Pseudoloma neurophilia: Progression of Infection and Transmission Characteristics of a Microsporidian Parasite in a Model Vertebrate, Danio rerio

Abstract approved:______Michael L. Kent

The microsporidian parasite, Pseudoloma neurophilia, is the most commonly diagnosed infectious disease in laboratory populations of the zebrafish, Danio rerio. Infections by P. neurophilia are generally subclinical, however, they can become acute either incidentally or due to experimental immune suppression. Non-protocol induced variation can confound results in laboratory experiments using such fish. As a result, there has been growing interest in sensitive diagnostic assays for P. neurophilia and demand for P. neurophilia specific-pathogen free zebrafish lines among the zebrafish research community. The high prevalence of P. neurophilia in zebrafish

provides the opportunity to investigate the progression of infection and transmission

characteristics of a microsporidian parasite in a well-developed model vertebrate host

species. I developed a real-time PCR-based assay combined with the use of sonication to improve spore disruption which has a sensitivity that is 10-100 times more sensitive than a previously published conventional PCR-based assay. The microsporidium infects ovaries and eggs, thus, I developed a sampling method for the testing of water from spawning fish and demonstrated the utility of testing spawn water, eggs, and sperm for the non-lethal detection of P. neurophilia in adult fish. The presence of P. neurophilia in spawn water and eggs from infected adults provided the initial evidence of vertical transmission of P. neurophilia in zebrafish. Intraovum transmission of P. neurophilia was directly observed by histological screening of larval fish spawned from infected adults and by analyzing eggs using a

transmitted light dissecting microscope. The prevalence of intraovum transmission was determined by screening surface-decontaminated eggs from 27 paired spawns using the real-time PCR based assay. Intraovum transmission was detected in 4 of the 27 spawns and the prevalence of intraovum P. neurophilia in the eggs from these spawns was determined to be approximately 1%. Parasite DNA was also detected in the spawning water from 11 of the 27 spawns, highlighting the potential for extraovum transmission of P. neurophilia. I investigated the early stages of P. neurophilia infection in experimentally infected larval zebrafish. This was accomplished using a combination of standard hematoxylin and eosin stain, the Luna stain, and an in situ hybridization probe specific to the small-subunit ribosomal DNA gene of P. neurophilia, which I developed. At 12 hours post exposure, P. neurophilia was mainly visualized as intact spores in the intestinal lumen, and proliferative stages developing in the epithelial cells of the anterior intestine and the pharynx, and within the hepatocytes of the liver. Proliferative (presporogonic) stages were visualized in these tissues and in the pancreas and kidney at 36 and 48 hours post exposure and in the spinal cord, eye, and skeletal muscle beginning

at 72 hours post exposure. The first spore stages of P. neurophilia were observed at 96

hours post exposure in the pharyngeal epithelium, liver, spinal cord and skeletal

muscle. The parasite was observed in the brain of larval fish at 120 hours post- exposure. The distribution of the early stages of P. neurophilia and the lack of mature spores until 96 hours post exposure suggests that the parasite gains access to organs distant from the initial site of entry, likely by penetrating the intestinal wall with the polar tubule, and that autoinfection does not occur at any detectable frequency during the initial stages of infection.

Pseudoloma neurophilia: Progression of Infection and Transmission Characteristics of a Microsporidian Parasite in a Model Vertebrate, Danio rerio

by Justin L. Sanders

A DISSERTATION

submitted to

Oregon State University

in partial fulfillment of

the requirements for the

degree of

Doctor of Philosophy

Presented September 3, 2013

Commencement June 2014

Doctor of Philosophy dissertation of Justin L. Sanders presented on September 3, 2013.

APPROVED:

Major Professor, representing Microbiology

Chair of the Department of Microbiology

Dean of the Graduate School

I understand that my dissertation will become part of the permanent collection of Oregon State University libraries. My signature below authorizes release of my dissertation to any reader upon request.

Justin L. Sanders, Author

ACKNOWLEDGEMENTS

There have been numerous people who have helped me in some way on this journey to whom I owe a huge debt of gratitude. First and foremost, I thank Dr. Michael Kent. His mentorship and guidance, as well as his infectious enthusiasm and optimism have made this work possible and he has provided me with a multitude of great opportunities. His encouragement to always "keep paddling" has proven invaluable in research, writing, surfing, and life. I also must thank my thesis committee members, Drs. Luiz Bermudez, Kathy O'Reilly, Michelle Steinauer, and Robert Tanguay, for their guidance, advice, and time. I must thank my former employers and mentors who encouraged me to embark on this journey, especially Sandi Fredrickson, Dr. Patty McVay, and Jerry Hurst, and my friends and colleagues at the Humboldt County Public Health Branch. This project would have been far more difficult if not for my training and experience as a public health microbiologist. To that I owe the excellent training staff at the California State Viral and Rickettsial Disease/Microbial Disease laboratories, and the

staff at the numerous public health laboratories in which I had the great opportunity to

train.

I want to thank the faculty and staff of the Department of Microbiology here at OSU for all of their help and time. I also want to thank Carrie Barton and Carrie Buchner of the Tanguay Laboratory for maintaining and providing the Pseudoloma-free fish which were used in the majority of the experiments for this project. Thanks also to the staff at the histology laboratory here at OSU, Kay Fisher, Misty Corbus and Renee Norred, for their excellent histotechnical assistance and the hundreds of high-quality slides which were prepared during the course of this project. A big thank you to the members of the Kent Laboratory, especially Virginia Watral, for putting up with me during these years and Dr. Trace Peterson for his fish pathology expertise, friendship, and hours of comic relief.

I also want to thank the many friends and colleagues I have made during my time here whose encouragement, collaborations, and discussions have helped me immensely throughout this project and will continue long afterward: Drs. Ann Cali, Peter Takvorian, Louis Weiss, Kevin Lafferty, Liz Didier, Nicolas Corradi and so many others. I am also grateful to Mark Francis and the folks at Aquaneering for sponsoring the fish disease section of the zebrafish husbandry session at the Aquaculture America meeting of the World Aquaculture Society. It has been very rewarding for me to give talks at this meeting and I have really enjoyed watching it grow every year. Major funding for this project came from the National Institutes of Health (NIH NCRR 5R24RR017386-02 and NIH NCRR P40 RR12546-03S1). A travel award from the College of Sciences assisted my travel to Tarrytown, NY to present at the International Workshop for Opportunistic Protists in 2012. Finally, I am also enormously grateful for my family, without whose constant love and support, this would not have been possible: My parents, Jim and Jeanne, my sister, Julie Gagner, who also provided excellent review and editing of this dissertation, and especially Tara, Gavin, and Maeve: this is for you.

CONTRIBUTION OF AUTHORS Virginia Watral assisted with data collection and experimental setup for the vertical transmission studies. Kerri Clarkson assisted with spawning fish and egg collection for the vertical transmission studies. Doctor Tracy Peterson provided assistance with the histological descriptions in the parasite progression study.

TABLE OF CONTENTS Page

Chapter 1. Introduction ...... 1

General Characteristics of the Microsporidia ...... 1

Impacts of Microsporidiosis ...... 1

Summary and Thesis Overview ...... 4

References ...... 5

Chapter 2. Microsporidiosis in zebrafish research facilities ...... 9

Abstract ...... 10

Introduction ...... 11

Current methods of detection ...... 13

Transmission ...... 15

Parasite surveillance...... 18

Facility design considerations ...... 19

Husbandry considerations ...... 21

Other considerations ...... 22

Going forward ...... 23

References ...... 27

Chapter 3. Development of a sensitive assay for the detection of Pseudoloma neurophilia in laboratory populations of the zebrafish Danio rerio ...... 30

Abstract ...... 31

TABLE OF CONTENTS (Continued) Page

Introduction ...... 32

Materials and methods ...... 34

Results ...... 41

Discussion ...... 43

References ...... 56

Chapter 4. Verification of intraovum transmission of a microsporidium of vertebrates: Pseudoloma neurophilia infecting the zebrafish, Danio rerio ...... 61

Abstract ...... 62

Introduction ...... 63

Results ...... 66

Discussion ...... 67

Methods...... 73

References ...... 83

Chapter 5. Early development and tissue distribution of Pseudoloma neurophilia in the zebrafish, Danio rerio ...... 88

Abstract ...... 89

Introduction ...... 90

Methods...... 91

Results ...... 93

Discussion ...... 95

References ...... 104

Chapter 6. The zebrafish as a model for microsporidiosis...... 106

Introduction ...... 107

TABLE OF CONTENTS (Continued) Page

History of zebrafish in research ...... 107

Advantages of the zebrafish model ...... 109

Development/characterization of zebrafish as a biomedical model ...... 114

Microsporidian infections of zebrafish ...... 116

Conclusion ...... 122

References ...... 136

Chapter 7. Summary and conclusions ...... 144

Bibliography ...... 147

Appendices ...... 164

Appendix A. Pleistophora hyphessobryconis (Microsporidia) infecting zebrafish (Danio rerio) in research facilities ...... 165

Appendix B. Ichthyosporidium weissii n. sp. (Microsporidia) infecting the Arrow Goby

(Clevelandia ios) ...... 190

LIST OF FIGURES

Figure Page

Figure 2.1. Wet mounts of microsporidian spores from zebrafish ...... 25

Figure 2.2. Histological sections of ovarian, intestinal and kidney infections of Pseudoloma neurophilia from zebrafish ...... 26

Fig. 3.1. Partial ssrDNA sequence alignment of Pseudoloma neurophilia ...... 49

Fig. 3.2. Pseudoloma neurophilia spore sonication time study ...... 51

Fig. 3.3. Pseudoloma neurophilia spore extraction method comparison by quantification cycle (Cq)...... 52

Figure 4.1. Spores of Pseudoloma neurophilia in Luna-stained histological sections of progeny of infected zebrafish, Danio rerio...... 77

Figure 4.2. Spores of Pseudoloma neurophilia in developing embryo of zebrafish, Danio rerio...... 78

Figure 4.3. Modes of transmission of Pseudoloma neurophilia in the zebrafish,

Danio rerio...... 79

Fig 5.1-5.4. Early stages of Pseudoloma neurophilia infection in the gastrointestinal

tissues of larval zebrafish at 12-72 hours post-exposure ...... 100

Fig. 5.5-5.8. Early stages of Pseudoloma neurophilia infection in extraintestinal organs of larval zebrafish ...... 101

Fig. 5.9-5.18. Early stages of Pseudoloma neurophilia in neural tissues of larval zebrafish ...... 102

Figure 6.1. Histological section of a 7 day post-fertilization larval zebrafish...... 124

Figure 6.2. Developing zebrafish embryo at 48 hours post-fertization with two opaque aggregates of Pseudoloma neurophilia ...... 125

Figure 6.3. Histological section of an adult zebrafish showing aggregates of spores of P. neurophilia in the spinal cord ...... 126

LIST OF FIGURES (Continued)

Figure Page

Figure 6.4 Adult zebrafish, Danio rerio, infected with the microsporidian Pseudoloma neurophilia ...... 127

Figure 6.5. Histological section of an adult zebrafish with focal chronic inflammation and myodegeneration due to Pseudoloma neurophilia infection ...... 128

Figure 6.6. High magnification of a histological section of an adult zebrafish with chronic myositis caused by Pseudoloma neurophilia infection ...... 129

Figure 6.7. Wet mount of several aggregates of Pseudoloma neurophilia spores obtained from the hindbrain of an infected adult fish ...... 130

Figure 6.8. Histological section of a larval zebrafish (7 days post fertilization) with developing stages of Pseudoloma neurophilia present in the liver visualized by in situ hybridization ...... 131

Figure 6.9. Diagram illustrating the modes of transmission observed in Pseudoloma neurophilia infections in the zebrafish, Danio rerio...... 132

Figure 6.10. Histological section of an adult female zebrafish with mature spores and presporogonic stages of Pseudoloma neurophilia observed within a late-stage,

vitellogenic oocyte...... 133

Figure 6.11. Histological section of an adult zebrafish infected with Pleistophora hyphessobryconis...... 134

Figure 6.12. Adult zebrafish, Danio rerio, with skin removed infected with Pleistophora hyphessobryconis ...... 135

LIST OF TABLES

Table Page

Table 3.1. Experimental setup for two extraction comparison trials ...... 53

Table 3.2. Pseudoloma neurophilia spiked sample results from three separate trials ...... 54

Table 4.1. Detection of Pseudoloma neurophilia in egg pools obtained from paired spawning of Danio rerio adults and tested by qPCR and in spawning adult fish by histology ...... 80

Table 4.2. Dose response of larval zebrafish exposed to low numbers of Pseudoloma neurophilia ...... 82

Table 5.1. Results of histological examination of larval zebrafish at various times post exposure to Pseudoloma neurophilia...... 103

LIST OF APPENDIX FIGURES

Figure Page

Appendix Figure 1.1. Zebrafish infected with P. hyphessobryconis ...... 182

Appendix Figure 1.2. Wet mount of P. hyphessobryconis spores ...... 183

Appendix Figure 1.3. Histological sections of zebrafish infected with P. hyphessobryconis...... 184

Appendix Figure 1.4. Phylogenetic tree of Pleistophora hyphessobryconis and related microsporidia based on small subunit rRNA gene sequences...... 185

Appendix Figures 2.1,2.2. Macroscopic and low magnification microscopic images of gonadal lesion in female arrow goby, Clevelandia ios ...... 203

Appendix Figures 2.3-2.8. Light micrographs of spores of Ichthyosporidium weissii n. sp ...... 204

Appendix Figures 2.9-2.14. Histological sections of gonadal tissue from female arrow gobies, Clevelandia ios, infected with the microsporidium, Ichthyosporidium weissii n. sp...... 205

Appendix Figures 2.15-2.20. Transmission electron microscopy of spores and

developmental stages of Ichthyosporidium weissii n. sp...... 206

Appendix Figure 2.21. Hematoxylin and eosin-stained histological section of skeletal muscle of an arrow goby, Clevelandia ios, infected with Kabatana sp., a myocyte infecting microsporidian ...... 207

Appendix Figure 2.22. Phylogenetic tree of microsporidian small subunit (SSU) rRNA gene sequences inferred by Bayesian analysis...... 208

Chapter 1. Introduction General Characteristics of the Microsporidia The Microsporidia are a phylum of obligate intracellular eukaryotic parasites with species infecting virtually all animal phyla. The first description of a microsporidium in the literature was written in 1857 by Wilhelm von Nägeli (1857). At the time, the silk industry in Europe was greatly impacted by a disease known as pébrine, or pepper disease, a lethal disease of silkworms, so named by the characteristic black mottled appearance of infected silkworms. Nägeli observed particles in silkworms affected by pébrine and named the particles Nosema bombycis. The pathogenic nature of the Microsporidia was later characterized by Louis Pasteur who described the causative agent of pébrine and outlined methods by which to prevent it. Subsequently, numerous important diseases of insects have been attributed to microsporidia and they have even been investigated as possible agents of biological control of pest insects. Microsporidia are characterized by their small size, the production of an environmentally resistant spore which is the infectious stage of the parasite, and the polar tubule which is the infectious apparatus of the parasite. The spore gets its rigidity from a thick endospore layer composed of chitin. The rigidity of the spore is important not only for the protection of the organism in the environment, but also to provide the turgor by which the spore infects a host cell. Upon sensing the appropriate environmental cue, which varies by microsporidian species, osmotic pressure within the spore quickly increases by the splitting of trehalose into glucose. This pressure eventually builds to the point that the polar tubule is explosively everted, piercing the cell membrane of the host cell and injecting the sporoplasm of the parasite into the cytoplasm of the host cell. Impacts of Microsporidiosis In addition to insects, microsporidia infect a broad range of aquatic organisms including crustaceans, amphipods, and freshwater, saltwater and anadromous fishes. The impacts of microsporidian parasites on fish in aquaculture, wild populations, and research have been documented on several occasions (Lom and Dyková, 1992; Shaw and Kent, 1999), and microsporidian species belonging to some 18 genera have been described in 2

fishes (Lom, 2002; Lom and Nilsen, 2003). Most of these infections appear to be chronic with minimal host mortality. Infections by some species, however, can cause profound economic impact on wild fish and aquaculture hosts in terms of mortality and commercial quality of fish. Several of these microsporidia have been shown to impact fish either by directly killing the host or indirectly by reducing fecundity (Ramsay et al., 2009; Wiklund et al., 1996) or decreasing the commercial quality of farmed fish (Shaw and Kent, 1999). In humans, several species are often associated with opportunistic infections of immunocompromised hosts, especially AIDS patients (Didier, 2005; Didier et al., 1991; Shadduck and Greeley, 1989), however, immunocompetent individuals can also be infected (Didier, 2005; Wanke et al., 1996; Weber et al., 1994). These infections are most often characterized by self-limiting, chronic diarrhea (Müller et al., 2001; Wichro et al., 2005), however recent studies have shown the presence of the microsporidia, Encephalitozoon spp. and Enterocytozoon bieneusi, in asymptomatic individuals, several of whom were actively shedding parasites in stool (Sak et al., 2011a, 2011b). These recent findings along with other studies showing the presence of human-pathogenic microsporidia in several water sources including tertiary sewage effluent (i.e., treated by

mixed medium filtration and chlorination), surface water, and groundwater (Dowd et al.,

1998) suggest that the exposure of humans to microsporidia may be more widespread than previously thought and has resulted in the placement of Microsporidia on the National Institutes of Health (NIH) Category B list of biodefense pathogens and the Environmental Protection Agency (EPA) microbial contaminant candidates list of concern for waterborne contaminants. The ubiquity of the Microsporidia also makes them important natural pathogens of laboratory animals (Baker, 1998, 2003; de Kinkelin, 1980; Troemel et al., 2008). In addition to the direct impacts of infectious disease on laboratory animals (e.g. on population and overall health of the animals), experiments using animals infected with a pathogen such as a microsporidian parasite can have significant non-protocol induced variation as a result of the infection. For example, subtle changes in host-physiology such

as immune modulation in response to microsporidian infection can confound results obtained from experiments examining immune development and function (Baker, 2003). Also, experimental immunosuppression of laboratory animals which harbor chronic microsporidian infections can lead to the unexpected death of these animals (Baker, 2003; Horváth et al., 1999; Sanders et al., 2010) While numerous microsporidia are known to infect mammals, many more have been recorded from fishes (Lom, 2002; Lom and Nilsen, 2003). The zebrafish, Danio rerio, is an important laboratory model for toxicology, developmental biology, cancer, and infectious disease research. It has a well-characterized immune system (Traver et al., 2003), and the complete genome has been sequenced with the genetic map showing an overall highly conserved synteny with the human genome (Postlethwait et al., 1998). Zebrafish have been used in numerous in vivo experiments using various bacterial and viral pathogens (Allen and Neely, 2010; Dooley, 2000; Ingham, 2009) to investigate host-pathogen interactions. The results of several infectious disease studies using zebrafish have shown a number of immune response pathways which are analogous to those of humans and other mammals (Meeker and Trede, 2008; Meijer and Spaink,

2011).

Due to the increased use of fish in research, chronic infections of zebrafish by

microsporidia can have confounding impact on experimental results using such fish (Kent et al., 2012). One microsporidium, Pseudoloma neurophilia was detected in over 74% of laboratory zebrafish colonies examined through the Zebrafish International Resource Center diagnostic service in 2010 (Murray et al., 2011), making it the most common pathogen in laboratory zebrafish. A few impacts of P. neurophilia infections on zebrafish have been documented. For example, fish infected with P. neurophilia have a higher incidence of emaciation and vertebral malformations (Matthews et al., 2001). Infection by P. neurophilia has also been demonstrated to be correlated with reduced growth in certain lines of fish, and with a significantly reduced fecundity of infected fish in the presence of concomitant stress (Ramsay et al., 2009).

The high prevalence of a microsporidian parasite in laboratory zebrafish populations, Pseudoloma neurophilia, provides an ideal opportunity to learn more about the fundamental characteristics of microsporidian infections using a well-characterized vertebrate model organism. Summary and Thesis Overview The subject of this dissertation is a description of the transmission characteristics and early stages of infection of the microsporidian parasite, Pseudoloma neurophilia, in a model vertebrate host, the zebrafish, Danio rerio. My objectives are twofold: 1. To determine the potential routes of transmission of P. neurophilia in D. rerio, namely to answer the question of whether vertical transmission occurs, and 2. To describe the initial stages of development and progression of P. neurophilia infection in larval zebrafish. Chapter 2 provides an overview of the body of knowledge regarding microsporidiosis of zebrafish in laboratory colonies and serves as background information for this thesis. This chapter focuses on the two species known to occur in laboratory zebrafish colonies, Pseudoloma neurophilia and Pleistophora hyphessobryconis, and describes the issues of microsporidiosis in laboratory zebrafish, methods of diagnosis, modes of transmission, and approaches to control of these

pathogens. This chapter was published as a review article in an issue of the Institute for

Laboratory Animal Research Journal (Sanders et al., 2012). In Chapter 3, I describe the development of a real-time PCR assay for the detection of P. neurophilia and of a sampling method to detect P. neurophilia in zebrafish tissues, spawning water and eggs. When applied to the testing of spawn water and eggs from a group of P. neurophilia infected adult fish, this method provides some of the first solid evidence for the vertical transmission of P. neurophilia. The methods in Chapter 3 provided the tools to investigate the vertical transmission of P. neurophilia in zebrafish as is described in Chapter 4. This assay has been adopted in numerous facilities to screen zebrafish for the presence of P. neurophilia. In Chapter 4, I characterize the vertical transmission of P. neurophilia in zebrafish. This study is the first to document the direct observation of intraovum

transmission of a microsporidian in a vertebrate species. In addition to the direct observation of vertical transmission by microscopy, the prevalence of intraovum transmission is determined using the real-time PCR and sampling methodology described in Chapter 3. In Chapter 5, the early stages and progression of P. neurophilia in larval zebrafish are described. Early proliferative stages of microsporidians are generally not visible using standard histological methods. By using a combination of DNA in situ hybridization and histology, I have been able to observe the tissues and patterns of infection in the initial stages of P. neurophilia infection. Chapter 6 summarizes many of the findings from these earlier chapters in the context of the development of the zebrafish as a model for microsporidiosis. A comprehensive review of the history and use of zebrafish as a model organism is provided. This chapter is being published in the second edition of “The Microsporidia and Microsporidiosis” that is currently in press. In the appendix, I include two additional publications on fish microsporidia contributed during the course of my thesis research. One describes the discovery of another microsporidium in zebrafish from laboratory colonies, Pleistophora

hyphessobryconis. The second is the description of a novel microsporidian species,

Ichthyosporidium weissii, infecting the arrow goby. This publication is notable because the lesions caused by the species described had been initially misdiagnosed as neoplasia, resulting in some alarm due to the possibility of contaminants in Moro Bay, California causing tumors in fish. REFERENCES Allen, J.P., and Neely, M.N. (2010). Trolling for the ideal model host: zebrafish take the bait. Future Microbiol. 5, 563–569.

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Chapter 2

Microsporidiosis in zebrafish research facilities

Justin L. Sanders1, Virginia Watral1, Michael L. Kent1,2

ILAR Journal Oxford University Press, Oxford, OX2 6DP, UK Issue 53: pp. 106–113

1Department of Microbiology. 220 Nash Hall, Oregon State University, Corvallis, OR 97331-3404 2Department of Biomedical Sciences, Oregon State University, Corvallis, OR 97331

ABSTRACT Pseudoloma neurophilia (Microsporidia) is the most common pathogen detected in zebrafish (Danio rerio) from research facilities. The parasite infects the central nervous system and muscle, and may be associated with emaciation and skeletal deformities. However, many fish exhibit subclinical infections. Another microsporidium, Pleistophora hyphessobryconis, has recently been detected in a few zebrafish facilities. Here we review the methods for diagnosis and detection, modes of transmission, and approaches used to control microsporidia in zebrafish, focusing on P. neurophilia. The parasite can be readily transmitted by feeding spores or infected tissues, and we showed that cohabitation with infected fish is also an effective means of transmission. Spores are released from live fish at various points, including the urine, feces, and sex products during spawning. Indeed, P. neurophilia infects both the eggs and ovarian tissues, where we found concentrations ranging from (12,000 – 88,000 spores/ovary). Hence, various lines of evidence support the conclusion that maternal transmission is a route of infection: spores are numerous in ovaries and developing follicles in infected females, spores are present in spawned eggs and water from spawning tanks based on PCR tests, and larvae are very susceptible to the infection. Furthermore, egg surface disinfectants presently

used in zebrafish laboratories are ineffective against microsporidian spores. At this time,

the most effective method for prevention of these parasites is avoidance.

INTRODUCTION The dramatic increase in the use of zebrafish (Danio rerio) in biomedical research has led to a corresponding increased interest in the diseases affecting this important biological model. Many of the laboratory animal health and pathogen control principles developed for mice and rats are applicable to aquatic laboratory animals such as the zebrafish, however, there are special considerations in working with aquatic animals. Kent et al. (2009) provided a general review of the control of diseases in fish research colonies. The present review focuses specifically on the transmission and control of microsporidia in zebrafish facilities. We emphasize particularly Pseudoloma neurophilia as this microsporidium is very common in these fish (Murray et al., 2011), and provide a discussion on Pleistophora hyphessobryconis, which was recently detected in a few facilities (Sanders et al., 2010). Microsporidia Microsporidia are obligate intracellular eukaryotic parasites with species infecting virtually all animal phyla. They have a relatively simple life cycle, consisting of two general developmental stages; mergony and sporogony. Meronts multiply inside the infected host cell, eventually forming sporonts and then spores, which are ultimately

released from the host and transmit the infection. The infectious spore stage has a thick,

chitinous endospore, making it extremely resistant to environmental stress and lysis, allowing the organism to maintain viability for extended periods in the aquatic environment (Shaw et al., 2000). Additionally, microsporidia are generally resistant to many standard forms of surface decontamination used for fish eggs such as chlorine and iodophores, complicating the control of these pathogens. Microsporidia are common pathogens of numerous aquatic organisms including crustaceans, amphipods, and members from some 18 genera of these parasites have been described in fishes (Lom, 2002; Lom and Nilsen, 2003). The impacts of microsporidian infections on fish populations in the wild, aquaculture, and laboratory have been documented in numerous cases (reviewed in Shaw and Kent 1999). These often focus on the more acute effects of microsporidian disease such as mortality, however, most

microsporidian species infecting aquatic animals result in chronic diseases with minimal associated host mortality (Murray et al., 2011). Pseudoloma neurophilia Pseudoloma neurophilia was first reported by de Kinkelin (1980) in fish purchased from a pet store for use in toxicological studies. The parasite was further described and assigned to a new genus, Pseudoloma neurophilia, by Matthews et al (Matthews et al., 2001). Pseudoloma neurophilia is the most commonly observed microsporidian parasite of zebrafish. For example, the infection was detected in greater in 74% of the facilities examined through the Zebrafish International Resource Center (ZIRC) diagnostic service in 2010 (Murray et al., 2011). It generally causes chronic infections in zebrafish with clinical signs ranging from emaciation and obvious spinal deformities (lordosis, scoliosis) to subclinical infections exhibiting no outward signs of disease (Matthews et al., 2001). As with other animals used in research, experiments utilizing zebrafish with these infections may be subject to non-experimental variation, potentially confounding results as has been described in laboratory colonies of rabbits and mice infected with the microsporidian parasite, Encephalitozoon cuniculi (Baker,

2003). Furthermore, infected fish without overt clinical disease have been shown to have

reduced fecundity and size (Ramsay et al., 2009).

Pleistophora hyphessobryconis The muscle-infecting microsporidium, Pleistophora hyphessobryconis, has also been observed and described in laboratory populations of zebrafish (Sanders et al., 2010). Commonly known as "neon tetra disease" for its type host, the neon tetra, Paracheirodon innesi, this parasite is very common in the aquarium trade, often resulting in considerable mortality. This microsporidium has been described in a broad range of fish hosts, and has been reported from many species of aquarium fishes in several families, including Danio rerio and D. nigrofasciatus (Steffens, 1962). Similar to Pseudoloma neurophilia, P. hyphessobryconis can also be harbored by otherwise healthy appearing fish, which may show clinical signs of the infection or mortality after experiencing experimental or incidental immunosuppression (Sanders et al., 2010). The presence of P.

hyphessobryconis infections in laboratory zebrafish colonies highlights the importance of obtaining fish used in research from reputable sources and also illustrates the potential for introduction of otherwise novel microsporidia with a broad host range to new hosts. CURRENT METHODS OF DETECTION External Indicators of Infection External indications of zebrafish infected by P. neurophilia include reduced growth, emaciation, spinal deformation (e.g. lordosis, scoliosis), or low-level mortalities with no grossly-visible lesions. Typically, indicators of infection and mortality become apparent only after a stress event (Ramsay et al., 2009), such as crowding or shipping. These general clinical presentations are not pathognomonic for P. neurophilia, making external examination of fish alone of little use in the diagnosis of this infection. The skeletal muscle infecting microsporidium, Pleistophora hyphessobryconis, can also be harbored by otherwise healthy appearing fish. Similar to fish infected with P. neurophilia, immunosuppression by various means can result in acute infection, with affected fish displaying large, depigmented regions localized around the dorsal fin. Fish presenting severe signs of P. hyphessobryconis eventually die from the infection.

Microscopy

Microsporidian spores can often be seen in wet mount preparations from infected

tissues. They are discernable by their generally refractile appearance and characteristic posterior vacuole. In suspected cases of infection by P. neurophilia, posterior brain and spinal cord tissue can be examined by wet mount for the presence of spores which are about 3 by 5 µm in size and pyriform in shape (Figure 2.1a). Wet mount preparations of tissue from opaque lesions present in the skeletal muscle can be examined for the presence of P. hyphessobryconis spores, which are 4 by 6-7 µm in size, also pyriform in shape and possess a very prominent posterior vacuole (Figure 2.1b). In general, microsporidian spores can be readily detected in standard hematoxylin and eosin (H&E) stained tissue sections when they occur in aggregates. However, in light infections, when only single spores are present within areas of inflammation, detection by H&E is difficult. Microsporidian spores appear Gram positive in Gram stains (Figures

2.2a, 2.2b, 2.2c, 2.2g) and are generally acid fast in various acid-fast staining methods (Figure 2.2e). The acid fast character of the spores can be variable depending upon the amount of decolorization. In cases where microsporidian infection is suspected, special stains such as the Luna stain or periodic acid Schiff (PAS) can greatly increase the visibility of spores allowing greater sensitivity of detection by histology (Peterson et al., 2011). Chitin specific fluorescent stains such as Fungi-Fluor (Polysciences, Warrington, PA) also increase the sensitivity of spore detection by histology but require the use of a fluorescence microscope (Kent and Bishop-Stewart, 2003). With Pseudoloma neurophilia, large aggregates of spores are primarily found in the neural tissue of the posterior brain and spinal cord. Smaller groups or individual spores can also be seen in the kidney, skeletal muscle, gut epithelium, and ovary (Kent and Bishop-Stewart, 2003), or within developing follicles (Figure 2.2). Spores of P. neurophilia released from aggregates within myocytes or peripheral nerves in the somatic muscle typically elicit a severe inflammatory reaction (Ramsay et al., 2009). In contrast, the muscle is the primary site of infection for Pleistophora hyphessobryconis. Massive infection by proliferative stages and spores occupy the myocyte, with inflammatory changes occurring after infections become so severe that the

myocytes rupture. Spores of this parasite can also be observed in the kidney, spleen,

intestine, and ovaries in heavier infections (Sanders et al., 2010). Molecular Diagnostics Conventional PCR (Murray et al., 2011; Whipps and Kent, 2006) and qPCR based (Sanders and Kent, 2011) assays targeting unique portions of the small subunit ribosomal DNA (ssrDNA) gene are available for testing of zebrafish tissues for P. neurophilia. The qPCR assay of Sanders and Kent, in combination with sonication, has also been applied to detect P. neurophilia ssrDNA in water, sperm, and eggs, providing a potential non-lethal assay for screening populations of fish for this parasite. As with most PCR-based assays, these tests are very sensitive and provide a relatively fast method of screening for the presence of P. neurophilia in zebrafish. No PCR-based assays currently

exist for the specific detection of P. hyphessobryconis, but this is a potential target for future studies. TRANSMISSION In order to control the spread of a pathogen in a population, it is important to understand its mode or modes of transmission. In general, microsporidia infecting fish are transmitted directly, presumably per os via ingestion of infected tissues or spores present in the water (Dyková and Woo, 1995; Shaw and Kent, 1999). The two microsporidia thus far described in zebrafish, P. neurophilia and P. hyphessobryconis, have been shown to infect fish by this method by experimental exposure (Kent and Bishop-Stewart, 2003; Sanders et al., 2010). Thus removal of dead and moribund fish would be expected to limit the potential exposure of tank mates to these two parasites. Murray and colleagues (2011) reported the spread of the parasite within a tank from < 6% to 77% prevalence over one year. They also showed that detritus from positive tanks placed in tanks containing parasite-free fish could spread the infection. We have found that live, infected fish transmit P. neurophilia by shedding it in the water, infecting recipient fish held in the same water but separated from each other by a screen

cage. Five flow through cohabitation tanks were set up using infected “donor” fish

segregated within a suspended breeding cage with a screen bottom placed in the same

tank with uninfected recipient zebrafish obtained from the P. neurophilia specific pathogen free colony housed at the Sinnhuber Aquatic Research Laboratory (SARL) at Oregon State University (Kent et al., 2011). One control tank was set up which consisted of both recipient and donor fish from the negative fish stock. After 2 months of cohabitation, donor fish were removed and posterior brains and spinal cords were examined by wet mount for the presence of P. neurophilia. The overall prevalence of P. neurophilia in the donor fish was 81% with no spores detected in the 10 negative control donor fish. Histological examination of the donor fish revealed that 3 experimental tanks, and the negative control tank, contained both male and female donor fish, while a fourth experimental tank contained all males except one immature female. After an additional 2 months, the recipient fish were euthanized and examined by histology to determine

infection status. Recipient fish from all positive tanks were infected, with an overall incidence of 66%. No infection was detected in the negative controls. Tank 4, which contained no sexually mature female donor fish, showed 57% incidence of infection. These results provide evidence that P. neurophilia is shed by live infected fish, and illustrate the route by which the parasite can spread throughout a population of fish in a single tank. This finding is consistent with other reports that Loma salmonae, a microsporidian parasite of salmonids, is similarly transmitted to tank mates by cohabitation (Ramsay et al., 2003; Shaw et al., 1998). The potential routes by which P. neurophilia may be transmitted by live, infected fish become apparent by observing the tissue distribution of the parasite. While P. neurophilia primarily targets neural and muscle tissue, we occasionally observe spores in the gut epithelium (Figure 2.2f) and in the kidney tubules (Figure 2.2g), each of these tissues providing a portal through which infectious spores can be shed into the water through feces or urine. Additionally, Kent and Bishop-Stewart (2003) reported the frequent occurrence of spores in the ovarian stroma (Figure 2.2b) and since that report we have also detected spores within developing follicles (Figures 2.2a, 2.2c, 2.2d, 2.2e), supporting maternal transmission during

spawning as another likely route of infection.

It is difficult to quantify microsporidian spores in histological sections, and thus

entire ovaries from females from nine separate infected populations were surveyed to more precisely determine the concentration of P. neurophilia (unpublished observations). Ovaries of 10 fish from each population were pooled, homogenized, and a sample of spores counted by hemocytometer. The average number of P. neurophilia spores seen was 44,000 per fish (range 12,000 – 88,000). Zebrafish frequently spawn spontaneously in aquaria, and hence release of eggs, ovarian fluids, and tissues at spawning provides an important potential route of horizontal transmission. However, the fact that recipient fish were positive from the tank in which donor fish had no sexually mature females suggests that spores are also released from infected fish by routes other than spawning. Observation of spores in the renal tubules and the intestinal epithelium (Figure 2.2f, 2.2g) supports this hypothesis.

Sex products not only provide an important source of infection to tank mates of the same age cohort, but also a source of infection to progeny by maternal transmission. Indeed, this route of infection has been reported for other microsporidia of fishes. The potential for maternal transmission, either transovum or transovarial, has been reported for Loma salmonae (Docker et al., 1997), and Ovipleistophora ovariae (Kent and Bishop- Stewart, 2003). Phelps & Goodwin (2008) provided the most conclusive evidence for vertical transmission of fish microsporidia, showing the presence of the DNA from Ovipleistophora ovariae within spawned eggs of the golden shiner Notropis chrysoleucas by qPCR. Further evidence for the maternal transmission of P. neurophilia was observed in the experiment described by Sanders and Kent (2011), where parasite DNA was detected in the eggs and water from a group spawn of infected zebrafish. We have tested the spawn water and eggs of several other groups of fish, and consistently found PCR positive water and eggs (unpublished observations). There are other experimental and observational lines of evidence that suggest maternal transmission of P. neurophilia, either transovarial (pseudovertical, outside of the egg or sperm) or transova (true vertical, within the egg or sperm). Evidence of true vertical transmission of P. neurophilia was observed in a follow-up experiment

performed from a laboratory study described by Ramsey and colleagues (2009). Six week

old zebrafish (AB strain), obtained from the ZIRC were experimentally exposed to P. neurophilia spores at 10,000 spores/fish. At 8 weeks post-exposure, six pairs of fish were separately spawned and the embryos reared in individual covered beakers in sterile water. Three pairs of unexposed fish were spawned separately as a negative control with the progeny reared under identical conditions. After spawning, all adult fish were processed for histology and slides were stained using the Kinyoun acid fast method to determine infection status and tissue distribution of the parasite (Ramsay et al., 2009). At 8 weeks post-hatch, juvenile fish were euthanized, viscera removed, and the remaining tissues (spinal cord, somatic muscle, head) were placed in pools of five fish and DNA extracted for PCR analysis using the method of Whipps and Kent (2006). Pseudoloma neurophilia was detected in 2 of 3 pools of fry from one spawning pair. Histological analysis of the

adult pairs showed the presence of microsporidian spores in the spinal cord, ovary, and most importantly in developing follicles of the spawning female (Fig 2.2e). As these fry were raised in isolation from the original spawning pair and the parasite was seen developing in eggs from the female, there is evidence that the infection was transmitted vertically, either by infection of the eggs prior to fertilization or by the exposure of the larval fish to spores present in high numbers in eggs which did not develop further. However, as P. neurophilia spores were also observed in the ovarian stroma, transovarial transmission (i.e. via spores outside of eggs) cannot be excluded. There is limited evidence currently available for the potential for maternal transmission of Pleistophora hyphessobryconis. Schäperclaus (1941) found infections in 8 day old neon tetras which had been derived from infected parents, suggesting the possibility of maternal transmission. We observed spores of this microsporidium in the ovarian tissue of infected females (Sanders et al., 2010), but no spores were seen in developing follicles in this study. The low prevalence of this parasite in laboratory zebrafish colonies would seem to minimize the importance of this mode of transmission for P. hyphessobryconis.

PARASITE SURVEILLANCE

Routine monitoring

Routine disease and pathogen monitoring is important not only in the control of microsporidian parasites, but also for the detection of other pathogens as well as in the monitoring of the overall health of the colony (Kent et al., 2009). It is only through routine monitoring of healthy, as well as moribund fish, that colony managers can detect potential health problems in fish. No serological tests are presently available for zebrafish. Histological analysis is the best overall method for routine health monitoring of zebrafish due to the ability to assess all tissues and to detect novel pathogens which would not be detected by specific PCR-based assays. Screening of fish in specific tanks by PCR to determine the presence or prevalence of P. neurophilia is also recommended, however, careful consideration of sample size is required to ensure the statistical relevance of these data (Kent et al., 2009, 2011).

Sentinel program The use of a sentinel program is a very effective means to monitor microsporidian infections in laboratory colonies. Exposing a population of known uninfected fish to the untreated effluent from other tanks on the system allows facility managers to assess the infection status of fish in the system on a large scale. For the monitoring of chronic microsporidian infections such as Pseudoloma neurophilia it is recommended that sentinel fish be held at least 3 months prior to sampling (Kent et al., 2009). The presence of P. neurophilia or other microsporidian parasites in the sentinel fish is an indication that infected fish are present somewhere in the facility. Ultraviolet sterilization is a common feature in recirculating water systems. It is useful to hold a sentinel population exposed to effluent post UV treatment in order to assess the efficacy of the filtration and disinfection of effluent water. FACILITY DESIGN CONSIDERATIONS Receiving fish into the facility/quarantine The practice of “eggs only” movement of fish between facilities has been successfully used for years in salmonid aquaculture to exclude pathogens from salmon

facilities (Kent and Kieser, 2003). It is recommended that fish received in a facility as

embryos be held in quarantine isolation and a subset examined before introduction into

the main facility. Also, if possible, the parents of these fish should be examined for pathogens that may be maternally transmitted (e.g., P. neurophilia). It is recommended that the quarantine area be physically separated from the main housing area, with restrictions on staff entering the main facility from the quarantine area. After determining that the brood stock is not infected with a microsporidian parasite, the progeny may be moved into the main facility. The short generation time of zebrafish facilitates this process greatly, allowing managers to bring adults into quarantine, spawn them, and then move only the progeny of those adults which are screened and determined to be microsporidian free into the main facility. This approach was used to establish a specific pathogen free (SPF) for P. neurophilia zebrafish laboratory at Oregon State University (Kent et al., 2011). Now two wild type lines of these fish are available to the research

community through the Sinnhuber Aquatic Research Laboratory at Oregon State University. Separation of tanks within the main facility The separation of tanks in the main facility is very important in the control of microsporidia. As microsporidian spores are transmitted by water and horizontally by infected fish, splashes and mixing of fish in tanks may result in the spread of these parasites throughout the facility. In fact, we have observed the spread of P. neurophilia from a single tank of infected fish to other fish in separate tanks housed in the same unit in which the effluent water was discharged into an open tray and frequently splashed (unpublished observations). We have also seen P. hyphessobryconis transmitted in a similar way to fish housed on the same rack as infected fish (Sanders et al., 2010). The transmission of another aquatic parasite, Ichthyophthirius multifiliis, between tanks via aerosolization of water in a laboratory has also been demonstrated (Wooster et al., 2001). Thus, covering of tanks and minimizing splashing of effluent is key to controlling the spread of microsporidiosis as is the isolation of tanks with known infected fish from those which are microsporidian free or of unknown infection status.

UV sterilization of water in recirculating systems

Ultraviolet (UV) sterilization of municipal drinking water has been used for

several years to inactivate protozoan pathogens such as Cryptosporidium and Giardia. These systems have also been shown to be effective in the inactivation of microsporidian parasites of human health concern such as Encephalitozoon intestinalis at a dose of 6 mJ/cm2 (Huffman et al., 2002). The effectiveness of UV sterilization is highly dependent upon proper prefiltration of incoming water to remove particulates, cleaning the quartz sheath that the UV bulb is inserted into, and the replacement of UV bulbs at regular intervals. As stated previously, it is important to maintain a group of sentinel fish downstream of the UV treatment in order to assess its efficacy.

HUSBANDRY CONSIDERATIONS Egg disinfection The purpose of egg disinfection is to kill pathogens which are present on the surface of the eggs, preventing their spread to progeny and potentially other fish in the facility. This method had been successful in the control of many pathogens in salmon aquaculture (Kent and Kieser, 2003). For zebrafish eggs, bath treatment with 25 to 50 ppm sodium hyphochlorite for 10 min is generally the method recommended for disinfection (Harper and Lawrence, 2010). Unfortunately, this level of bleach is ineffective at killing P. neurophilia (Ferguson et al., 2007). A similar situation can be seen with the disinfection procedures for salmonid eggs, in which the iodine treatments used were shown to be ineffective at eliminating 100% of spores of Loma salmonae, even at very high levels of iodine (Shaw et al., 1999). Therefore, microsporidian spores are highly resistant to current methods of surface sterilization of eggs, and these methods cannot be relied upon to eliminate P. neurophilia or other microsporidia from a population, nor can it be relied upon to effectively prevent the spread of microsporidian parasites between fish colonies. Further compounding this problem is the potential for transmission of the parasite within eggs. Transovum (true vertical transmission) of this

parasite would prevent the efficacy of any surface decontamination of eggs for P.

neurophilia, thus requiring careful screening of fish and the use of SPF fish stocks to prevent the spread or introduction of the parasite. Screening of sperm, eggs, larval fish Current molecular diagnostic methods can easily be applied to the testing of eggs, sperm and larval fish. In fact, the method of Whipps and Kent (2006) was used to screen eggs and larval fish in the development of a P. neurophilia specific-pathogen free zebrafish colony at Oregon State University (Kent et al. 2011). The qPCR method of Sanders and Kent (2011) was shown to be effective in testing sperm and eggs with a sensitivity of 10 spores per µl and 2 spores per egg, respectively. The cryopreservation of zebrafish sperm presents a special problem for preventing the spread of microsporidians. While P. neurophilia has not been seen in the testes of fish (Murray et al. 2011), there is

the potential for contamination of sperm from the kidneys or gut of the fish during manual stripping. Further compounding this problem is the potential for survival of the parasite during cryopreservation. While the ability of P. neurophilia to survive during cryopreservation is unknown, Nucleospora salmonis, a microsporidian parasite infecting salmonids, is maintained for long periods by cryopreservation in tissue culture (Wongtavatchai et al., 1994). Also, cryopreserved spores of mammalian microsporidia, which are viable, are readily available from the American Type Culture Collection, Manassas, VA. Disinfection of equipment The resistance of infectious microsporidian spores to environmental conditions requires the use of appropriate disinfection procedures to control the spread of these pathogens. Chlorine is commonly used to disinfect tanks and other equipment in zebrafish facilities. Ferguson et al (2007) found that 100 ppm chlorine (pH 7) effectively kills > 95% of P. neurophilia spores. Unfortunately, this is lethal for embryos and this is not suitable for egg disinfection. We are not aware of any studies which specifically test the efficacy of chlorine on Pleistophora hyphessobryconis, but it is likely that it would be killed at similar concentrations.

OTHER CONSIDERATIONS

Several zebrafish lines which are specific pathogen free (SPF) for Pseudoloma neurophilia have been developed at the colony housed at the Sinnhuber Aquatic Research Laboratory (SARL) (Kent et al., 2011). The development of these SPF lines was facilitated by the construction of a new fish facility which enabled the introduction of fish only after they were determined to be free of P. neurophilia. These fish are rigorously screened in order to maintain their SPF status. Obviously, the control of this parasite in existing facilities is much more complex and requires systematic screening and isolation of zebrafish with known infections in order to eliminate or reduce the presence of P. neurophilia infections in the colony (Murray et al., 2011). There are currently no known treatments for microsporidiosis in zebrafish. However, Fumigillin DCH, an agent used to treat the microsporidium Nosema apis in

honey bees, has been shown to be effective for several microsporidia infecting fishes (Shaw and Kent, 1999). Albendazole and monensin also have some efficacy in the treatment of salmonids for infections by Loma salmonae (Speare et al., 1999, 2000). The use of these drugs on experimental fish, while potentially eliminating the pathogen, could also introduce other changes in the host, confounding research (Baker, 2003). Toxic effects of Fumigillin DCH have been observed in salmonids (Laurén et al., 1989), thus its utility would be limited to the treatment of fish not used as experimental animals (e.g., brood stock). Ultimately, the elimination of P. neurophilia from existing lines of zebrafish may require rederivation of those lines using the methods described by Kent et al. (2011). GOING FORWARD The chronic and often subclinical nature of P. neurophilia infections in zebrafish requires the use of rigorous screening methodologies in order to ascertain the true prevalence of this parasite in laboratory zebrafish colonies. Its continued presence in laboratory zebrafish facilities highlights the need for increased surveillance, implementation of biosecurity protocols, and further research into the transmission and

control of these pathogens. Future studies to determine the efficacy of decontamination

protocols, such as the dosage of UV required to inactivate spores of P. neurophilia in

water and the survivability of the parasite during cryopreservation are needed. Additionally, the potential for introduction of novel microsporidia to zebrafish facilities underscores the need to obtain fish from reputable suppliers who are able to provide a health history of the fish. We also strongly recommend that zebrafish be obtained from suppliers who do not maintain zebrafish with other aquarium fish species. As the treatment of zebrafish with antimicrosporidial drugs may exacerbate impacts on research outcomes, the only effective method of controlling P. neurophilia infections in zebrafish is identification and removal of infected fish and avoiding introduction of the parasite by proper quarantine and screening of incoming fish. Whereas methods to avoid the infection and SPF zebrafish are now available, we have seen little enthusiasm for using parasite-free zebrafish by some researchers. This is

often due to the perception that subclinical infections have little or no impact on research endpoints (Kent et al., 2012). Therefore, another research need is the demonstration of the specific physiological, immunological, molecular, behavioral, etc. changes associated with subclinical infections by this extremely common parasite of zebrafish. ACKNOWLEDGEMENTS This study was supported by grants from the National Institutes of Health (NIH NCRR 5R24RR017386-02 and NIH NCRR P40 RR12546-03S1). We would like to thank C. Kent for assistance in review of this manuscript.

Figure 2.1. Wet mounts of microsporidian spores from zebrafish. A) Aggregates of spores of Pseudoloma neurophilia, contained within sporophorous vesicles (arrow). B) Pleistophora hyphessobryconis from the skeletal muscle. Note prominent posterior vacuole in spores (arrow). Bar = 10 µm.

Figure 2.2. Histological sections of ovarian, intestinal and kidney infections of Pseudoloma neurophilia from zebrafish. Bar = 10 µm unless otherwise indicated. A) Gram-positive (blue) staining spores in follicles (arrows). Bar = 50 µm. B) Gram-positive spores (arrows) in stroma of ovary. C). Numerous, Gram positive spores in developing follicle. D) Developing follicle replete with spores. H&E. E) Spores within a developing follicle. Kinyoun acid fast stain. Note the faint acid-fast appearance of spores due to overdecolorization. F) Spores (arrow) in intestinal epithelium. G) H&E. Spores in renal tubule (arrow). Gram stain.

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Shaw, R., Kent, M., and Adamson, M. (1999). Iodophor treatment is not completely efficacious in preventing Loma salmonae (Microsporidia) transmission in experimentally challenged chinook salmon, Oncorhynchus tshawytscha (Walbaum) - Shaw - 2002 Journal of Fish Diseases - Wiley Online Library. J. Fish Dis. 22, 311–312.

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Shaw, R.W., Kent, M.L., Brown, A.M.V., Whipps, C.M., and Adamson, M.L. (2000). Experimental and natural host specificity of Loma salmonae (Microsporidia). Dis. Aquat. Organ. 40, 131–136.

Speare, D.J., Athanassopoulou, F., Daley, J., and Sanchez, J.G. (1999). A preliminary investigation of alternatives to fumagillin for the treatment of Loma salmonae infection in rainbow trout. J. Comp. Pathol. 121, 241–248.

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inhibition of xenoma-expression in trout challenged with Loma salmonae (Microspora). J. Fish Dis. 23, 231–233.

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Whipps, C.M., and Kent, M.L. (2006). Polymerase chain reaction detection of Pseudoloma neurophilia, a common microsporidian of zebrafish (Danio rerio) reared in research laboratories. J. Am. Assoc. Lab. Anim. Sci. 45, 36–39.

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Chapter 3

Development of a sensitive assay for the detection of Pseudoloma neurophilia in laboratory populations of the zebrafish Danio rerio

Justin L. Sanders1, Michael L. Kent1,2

Diseases of Aquatic Organisms Inter-Research Science Center, Oldendorf/Luhe, Germany Issue 96: pp. 145–156.

1Department of Microbiology. 220 Nash Hall, Oregon State University, Corvallis, OR 97331-3404 2Department of Biomedical Sciences, Oregon State University, Corvallis, OR 97331

ABSTRACT Zebrafish (Danio rerio) are an increasingly important biological model in many areas of research. Diseases of zebrafish, especially those resulting in chronic, sub lethal infections, are of great concern due to the potential for non-protocol induced variation. The microsporidium, Pseudoloma neurophilia, is a common parasite of laboratory zebrafish. Current methods for detection of this parasite require lethal sampling of fish, which is often undesirable with poorly spawning mutant lines and small populations. We present here an improved molecular based diagnostic assay using real-time PCR, and including sonication treatment prior to DNA extraction. Sensitivity was increased compared to the previously published conventional PCR-based assay based on a dilution experiment, showing that this new assay had the ability to detect parasite DNA in one log higher dilution than the conventional PCR-based assay, which did not include sonication. Comparisons of several DNA extraction methods were also performed to determine the method providing the maximum sensitivity. Sonication was found to be the most effective method for disrupting spores. Further, we demonstrate the application of this method for testing of water, eggs and sperm, providing a potential non-lethal method for detection of this parasite in zebrafish colonies with a sensitivity of 10 spores per liter, 2

spores per egg and 10 spores per µl of sperm, respectively.

INTRODUCTION The zebrafish, Danio rerio, is a widely used biological model in several fields including developmental biology, immunology, toxicology, infectious disease and cancer research (Amatruda et al., 2002; Dooley, 2000; Eisen, 1996; Hill et al., 2005). Laboratory colonies of zebrafish are typically composed of specialized mutant strains of fish possessing genotypes useful to specific areas of study, and hardier wild-type strains are used for breeding stock and for maintaining specific genotypes in a heterozygotic state (Westerfield, 2007). Maintenance and husbandry of many of the mutant strains is often difficult (Lawrence, 2007). Individual adult fish from these lines, therefore, are often in limited supply and may be extremely valuable. Whereas laboratory populations of zebrafish occasionally are affected by acute infectious diseases, the most important and more prevalent are chronic infections by Mycobacterium spp. and Pseudoloma neurophilia (Microsporidia) (Kent et al., 2009; Matthews, 2004). The latter has been detected in over 50% of the facilities that we have examined through the Zebrafish International Resource Center (ZIRC) diagnostic service. Pseudoloma neurophilia, a microsporidian parasite, generally causes chronic

infections in zebrafish with clinical signs ranging from obvious scoliotic changes and

emaciated appearance of fish, to subclinical infections exhibiting no outward signs of

disease (Matthews et al., 2001). As with other animals used in research, experiments utilizing zebrafish with these infections are subject to non-experimental variation, potentially confounding results as described in laboratory colonies of rabbits and mice infected with the microsporidian parasite, Encephalitozoon cuniculi (Baker, 1998). Furthermore, fish without overt clinical disease may have reduced fecundity associated with the infection (Ramsay et al., 2009). Microsporidia are obligate intracellular parasites with species infecting virtually all animal phyla. They have a relatively simple life cycle, consisting of two general developmental stages; mergony and sporogony. Meronts multiply inside the infected host cell, eventually forming sporonts and then spores, which are ultimately released from the host and transmit the infection. The infectious spore stage has a thick, chitinous outer

layer, making it extremely resistant to environmental stress and lysis, allowing the organism to maintain viability for extended periods in the aquatic environment (Shaw et al., 2000). While several assays exist for the detection of pathogens in other fish species using non-lethal sampling methods (Lindstrom et al., 2009; López-Vázquez et al., 2006; Miriam et al., 1997), these sampling methods are generally not applicable to the zebrafish due to its small size and difficulty in obtaining blood and other tissues. Because zebrafish can be housed in relatively small volume tanks, the screening of water in tanks and even effluent from flow-through systems seems a feasible method by which to detect this pathogen without requiring lethal sampling of adult fish. Also, zebrafish spawn frequently and thus sperm, eggs, etc. are readily available providing potential samples for testing. For Microsporidia infecting fish, the only viable stage found in water is the spore, as other stages of these obligate intracellular parasites would not survive long outside the host. Therefore, in contrast to presporogonic stages found in tissues, efficient disruption of spores is crucial for obtaining DNA in the development of sensitive assays that are

based on the detection of this stage. Disruption of the tough, chitinous spore stage of the

parasite requires special methods such as mechanical disruption by bead beating or

sonication (Docker et al., 1997; Dowd et al., 1998; Fournier et al., 2000; Graczyk et al., 2007; Hoffman et al., 2007; Müller et al., 1999; Phelps and Goodwin, 2007), the use of saccharolytic enzymes such as chitinase (Delarbre et al., 2001; Müller et al., 1999), or in vitro germination of the spore using chemical means such as hydrogen peroxide in combination with other purification methods (Higes et al., 2006). Additionally, the few genomic studies of microsporidian parasites have found a general pattern of small, streamlined genomes with very few gene copies (Williams et al., 2008). In fact, one genus has been found to possess a single copy of the small subunit ribosomal RNA gene (Cornman et al., 2009), highlighting the need to maximize spore disruption and DNA concentration to achieve a sensitive and practical method of detection as most PCR tests for microsporidia have been based on this gene (Brown and Kent, 2002; Joseph et al.,

2006; Zhu et al., 1993). Whipps and Kent (2006) developed a PCR test for P. neurophilia based on the detection of small subunit ribosomal RNA (ssrDNA) gene sequences, and it was capable of consistently detecting down to an estimated 10 spores/sample of brain and spinal cord tissue. Here we describe a new, more sensitive PCR test for P. neurophilia, using a real- time PCR platform. Our intent was to use this assay for screening water and sex products and an important focus of this study was determination of optimal extraction methods for detecting the spores in both media. We also included screening of water from spawning tanks, eggs and sperm for non-lethal testing as zebrafish spawn prodigiously and spores of P. neurophilia occur both within and outside eggs in ovaries (Kent and Bishop- Stewart, 2003; Kent et al., 2007). MATERIALS AND METHODS Pseudoloma neurophilia spore preparations. All Pseudoloma neurophilia spores used in this study were obtained from hind brain and anterior spinal cords from 40 known infected stock zebrafish that were euthanized by an overdose of tricaine methanesulfite (Argent Laboratories, Redmond,

WA). A modification of a previously described method (Ferguson et al. 2007) was used

to isolate spores. Briefly, hindbrain and spinal cord tissue were mixed with 5 ml sterile

phosphate buffered saline (PBS) and then homogenized by passing through successively smaller gauge needles and filtered through a 20 µm nylon mesh filter. This homogenate was then centrifuged through a 50% Percoll (Sigma-Aldrich, St. Louis, MO) gradient for 50 min at 1,200 g and further purified by washing in a 0.45 µm filter (Millipore, Billerica, MA) twice with (PBS) to minimize the number of presporogonic stages present, in order for results of the extraction studies to be based on spores only. The resulting spore suspension was eluted from the filter in PBS, quantified using a hemocytometer and then diluted in PBS as needed. Assay design. A Taqman-based PCR assay was used to measure P. neurophilia DNA using an ABI 7500 sequence detection system (ABI Biosystems, Foster City, CA). Primers were

designed to amplify the ssuRNA gene of P. neurophilia based on sequence available in the NCBI Genbank (accession number AF322654) and using the Primer-BLAST program also available online from the NCBI with the primer parameters set to search for a PCR product size from 70-125 bp, optimal primer melting temperature of 60°C, and BLAST parameters set to search for similarity in the NCBI non-redundant database (All GenBank + RefSeq Nucleotides + EMBL + DDBJ + PDB sequences (excluding HTGS0,1,2, EST, GSS, STS, PAT, WGS) ) for specificity. The forward primer Pn10F (5` GTAATCGCGGGCTCACTAAG 3`), and reverse primer Pn10R (5` GCTCGCTCAGCCAAATAAAC 3`) were selected on the basis of lack of identity to related microsporidia, annealing temperature and small amplicon size of 113 base pairs, from position 1175 to position 1288 on the ssuRNA gene. A 3` hydrolysis probe complementary to a 23 base pair section of the amplified region was designed using the sequence 5`- 6-carboxyfluorescein (FAM)-ACACACCGCCCGTCGTTATCGAA – 3` Black Hole Quencher 1 (BHQ1). All reactions were performed in 25 µl using 900 nM forward and reverse primers, 250 nM of hydrolysis probe, 1X TaqMan Universal PCR Master Mix (ABI) and 2 µl of sample extract. The real-time PCR was performed on an

ABI 7500 Sequence Detection system (ABI) using the following reaction conditions:

50oC for 2 min, 95oC for 10 min followed by 40 repetitions (95°C for 15s, 60°C for 1

minute). Data analysis was performed using the 7500 System SDS Software version 1.3.1 (ABI). Cross reactivity testing. Cross reactivity of the assay was performed using the PN10F/PN10R primer and the PN probe with DNA extracts obtained from spores of two fish microsporidian parasites that could potentially be encountered in a zebrafish research facility: Glugea anomala obtained from three-spined sticklebacks in a research colony and Pleistophora hyphessobryconis obtained from neon tetras from a commercial tropical fish vendor.

Evaluation of pretreatments and DNA extraction methods for spores. Sonication. Sonication was one of our primary pretreatment methods to test. Prior to comparing pretreatment and extraction methods in the two trials below, it was necessary to determine the optimal sonication time for the disruption of spores. Thus a time study was performed in which several suspensions of spores consisting of 1,000 spores in 100 µl of PBS were sonicated with a Branson Sonifier 250 (Branson, Danbury, CT) for various time points and 2 µl of the crude sonicate tested by the real-time PCR method described above. The time points tested were 15 – 900 sec (Figure 3.2) and all were run at 55 W at a frequency of 20 kHz. Samples were not cooled during the sonication period, but were immediately placed on ice after treatment. The probe was decontaminated between samples using 10% household bleach followed by rinsing with sterile water. Each time point was examined in triplicate. It was determined that 5 min consistently provided the lowest quantification cycle and thus this was used for the sonication method in the extraction comparison trials. Trial A. A preliminary experiment was performed to assess the efficacy of sonication and hydrogen peroxide pretreatments in conjunction with three DNA

extraction methods (Table 3.1). Trial A was performed using an initial 900 µl suspension

containing 30,000 spores in total. The suspension was divided into three, 300 µl aliquots and subjected to either no treatment or one of two pretreatments: Pretreatment with hydrogen peroxide. As numerous microsporidian spores have been shown to germinate in vitro in the presence of hydrogen peroxide (Keohane and Weiss, 1999) we attempted to germinate spores of P. neurophilia by adding 30 µl of 30% hydrogen peroxide to the spore sample for a final concentration of 9% and incubated at 30°C for 30 min. The suspension was centrifuged for 5 min at 14,000 x g in a tabletop centrifuge. The supernatant was removed, 300 µl of molecular grade water was added and the suspension was allowed to sit for 30 min prior to further treatment. Pretreatment with sonication. Samples were sonicated with a Branson Sonifier 250 (Branson, Danbury, CT) for 5 min and immediately placed on ice after treatment.

The probe was decontaminated between samples using 10% household bleach followed by rinsing with sterile water. Following pretreatment (none, sonication or hydrogen peroxide), each pretreated pool was then divided into six 50 µl aliquots and either tested directly or extracted in duplicate using the following methods before testing duplicate reactions by real-time PCR: QIAgen. The DNeasy Blood and Tissue Extraction kit (QIAgen, Inc., Valencia, CA) was used for this procedure following the manufacturer’s protocol for extraction of DNA from tissues, with the addition of a single overnight freeze-thaw of the spore suspension and an overnight proteinase K and lysis buffer digestion at 56°C. Samples were eluted in 100 µl of buffer supplied in the kit. MoBio. The UltraClean™ Microbial DNA Isolation kit (Mo Bio, Carlsbad, CA) employs heat, detergent lysis, and bead-beating using specialized bead tubes and a standard vortex mixer with an adapter plate. The manufacturer’s protocol was followed using the included reagents, bead tubes, and silica membrane centrifugal filter columns. Briefly, 50 µl of pretreated/non-pretreated spore suspensions were first added to detergent buffer and heated at 65°C for ten min after which they were placed in the bead

tubes and vortexed for another ten min. After vortexing, the DNA was bound to a silica

membrane centrifugation filter and cellular components were washed out with provided

wash buffer. Finally, DNA was eluted in 50 µl of tris buffer supplied in the kit. Trial B. Based on the results from Trial A, pretreatment with hydrogen peroxide was not used and sonication was investigated further in a second experiment which was performed using purified spore suspensions containing 1,000 spores each in 100 µl PBS. Here, pretreatment with bead beating and chitinase was tested, along with sonication (as described in Trial A) and no pretreatment (Table 3.1). QIAgen (with or without pretreatments) and MoBio extractions (according manufacturer instructions) were conducted as described in Trial A. Three spore suspensions were tested directly by real- time PCR with no pretreatment nor DNA extraction and the following extraction methods were compared using triplicate spore suspensions:

Bead beating. Spores were suspended in Buffer ATL with proteinase K, both from the QIAgen Blood and Tissue Extraction kit, and 500 mg of 0.5 µm glass beads (Sigma-Aldrich, St. Louis, MO) in 1.5 ml screw-cap tubes. The samples were then run on a bead beater (BioSpec, Bartlesville, OK) at high speed (4500 oscillations per minute) for 3 min after which they were incubated for two hours at 56°C and extracted following the QIAgen method described above. All samples were eluted in a final volume of 100 µl of buffer supplied in the QIAgen kit. Chitinase. Chitinase (0.4U, Sigma-Aldrich, St. Louis, MO) and 200 µl potassium phosphate buffer (200 mM, pH 6.0) was added to each sample which was then incubated at 37°C for 30 min. Proteinase K from the Dneasy kit was then added and the samples were processed using the QIAgen method per manufacturer’s protocol and eluted in a final volume of 100 µl of buffer supplied in the QIAgen kit. Statistical analysis. Levene’s test for equality of variances was performed on the Cq values obtained in Trial B. After determining that there were no significant differences between variances in the extraction methods (Levene’s test, p=0.73), a One- way Analysis of Variance (ANOVA) was performed. Multiple comparison with best procedure based on Hsu’s method (Kuehl, 2000) was performed to determine the

methods which provided the highest sensitivity. All analyses were performed using the

statistical package R (www.r-project.org). Based on results from the extraction method study, all further experiments employed sonication or sonication followed by QIAgen extraction. Detection limit of real-time PCR Spores in PBS. A spore suspension was prepared as described above and spore suspensions were made in triplicate using 1,000 spores in 100 µl PBS, 100 spores/ 100 µl PBS, and 10 spores/ 100 µl PBS. Each suspension was sonicated and 2 µl of the crude sonicate was directly analyzed using the real-time PCR method described above in order to obtain 20, 2, and 0.2 spores per real-time PCR reaction, respectively. Spores in Group Spawn Tank Water. The ability of the test to detect the parasite in spawn water was evaluated. Ten liters of fish system water was divided into 1 L flasks

and inoculated as follows: three 1 L aliquots with 10 spores, three 1 L aliquots with 50 spores, three 1 L aliquots with 100 spores, three 1 L aliquots with 500 spores and one 1 L aliquot remained as a negative control. Each spiked 1 L water sample was individually filtered through a 1.2 µm nitrocellulose filter (Millipore #RAWP04700) in a fritted glass filter holder (Millipore, Billerica, MA) using a vacuum pump at 300 mm Hg. After filtration, the filter was rolled up using sterile forceps and placed in a 1.5 ml conical tube screw-cap tube. The water filtration apparatus was washed and decontaminated with 10% household bleach followed by rinsing in sterile water between samples. One ml of acetone was added to each 1.5 ml conical tube with the nitrocellulose filter and vortexed for several seconds. The tubes were then centrifuged at maximum speed for 3 min (> 16,000 rcf). The acetone supernatant was carefully aspirated off of the pellet with a transfer pipette. This was repeated two times to ensure all dissolved nitrocellulose was removed from the sample. One ml of 100% ethanol was then added to each tube and the pellet was suspended by vortexing and centrifuged at 3,000 g for 5 min. The ethanol was again carefully aspirated and 5 ml of 70% ethanol was added to each

tube. Tubes were centrifuged for 5 min at 3,000 g and the ethanol was again aspirated.

The pellet was resuspended in 100 µl of phosphate buffered saline (PBS) and then was

sonicated for 5 min at 55W. After sonication, each sample was placed on ice and DNA was extracted using the QIAgen extraction protocol described above. All samples were eluted in a final volume of 200 µl of buffer supplied in the kit. Spores in Eggs. We conducted the following test to evaluate detection of spores associated with eggs. A total of 2,000 eggs were obtained from the SPF zebrafish colony at the Sinnhuber Aquatic Research Laboratory (SARL), Oregon State University and divided into four aliquots of 500 eggs each with PBS added to make a total volume of 1 ml. One aliquot was spiked with 100 spores, one with 1000 spores, one with 10,000 spores and the last aliquot had no spores (negative control). Each sample was then sonicated for 5 min at 55W and cooled on ice. After sonication, three 40 µl aliquots (equaling approximately 20 eggs based on the original volume) were taken from each

sample and DNA was extracted using the QIAgen blood and tissue protocol as described above. All samples were eluted in a final volume of 200 µl of buffer supplied in the kit. Spores in Sperm. Sperm was obtained from SPF zebrafish males from the colony at the SARL by squeezing to evaluate the test with this sex product. The sperm samples were pooled and divided into 5 µl aliquots in 100 µl of PBS and spiked, in triplicate, with 5 spores/sample, 50 spores/sample and 500 spores/sample. All samples were brought to a volume of 200 µl and were then sonicated and DNA was extracted using the QIAgen extraction described above. All samples were eluted in a final volume of 200 µl of buffer supplied in the kit. The spiked spawn water, eggs and sperm were run in triplicate on different days. Comparison with conventional PCR assay. We compared our new real-time PCR based assay, which includes sonication pretreatment, with the previously developed conventional PCR based test described by Whipps and Kent (2006), which does not include sonication. The hindbrain and spinal cords from six infected fish were removed and individually homogenized in 100 µl of sterile PBS. Then 50 µl of the resulting homogenate was sonicated for 5 min followed by purification using the QIAgen method

and the remaining 50 µl was extracted by the QIAgen method as described by Whipps

and Kent (2006). All DNA extracts were eluted in 100 µl of tris buffer and four serial

tenfold dilutions of each was made in sterile water. After extraction, all samples and dilutions were run in single reactions using real-time PCR as described here and the conventional method as described by Whipps and Kent (2006) with minor modification. Briefly, the primer pair Pn18S5F 5′ GAA AAT TAC CGG AGC CTG AAG TC 3′, and Pn18S5R, 5′ TTC CCT CTC TCT CCA AAT TTC GG 3′were used to amplify a 788 bp fragment of the ssrDNA of P. neurophilia using conventional PCR. The reaction was carried out in 25 µL volumes using the Platinum® PCR SuperMix (Invitrogen, Carlsbad, CA), 12.5 pmol of each primer and 2 µl of extracted DNA. Amplification was carried out on a PTC-200 thermocycler (MJ Research, Watertown, MA) with an initial denaturation at 94°C for 3 min followed by 35 cycles of 94°C for 30 s, 55°C for 30 s, 72°C for 60s, and a final extension step of 72°C for 7 min. Products were visualized on a 1.5% agarose

gel stained with SYBR® Safe DNA Gel Stain (Invitrogen, Carlsbad, CA). Results were reported as positive (P. neurophilia ssrDNA detected) or negative (no P. neurophilia ssrDNA detected) based on the presence or absence of a band corresponding with an approximate size of 788 base pairs. Group Spawn Experiment. The ability of the test to detect the parasite in sex products of infected fish was evaluated. Ten adult zebrafish were arbitrarily selected from a population determined to have a 10% prevalence of Pseudoloma neurophilia based on histological examination of a subsample of fish three months prior to the experiment. The fish were placed in a spawning tank with 10 l of system water overnight. The following day, the fish were collected and euthanized by an overdose of tricaine methanesulfonate (Argent Laboratories, Redmond, WA). Brains and spinal tissue were collected using sterile instruments between individuals, placed in separate 1.5 ml tubes and extracted using the standard QIAgen protocol with an overnight digestion at 56°C. Water was filtered in 1 l aliquots using the method described previously. Filters were dissolved, sonicated and extracted as described previously. Eggs were pooled, sonicated and five 40µl aliquots were extracted as described previously. All samples were tested in single reactions using the real-time PCR method.

RESULTS

Cross Reactivity Primer-BLAST search of the 20 base pair forward primer, PN10F and the 20 base pair reverse primer, PN10R returned AF322654.1 Pseudoloma neurophilia small subunit ribosomal RNA gene, partial sequence; internal transcribed spacer, complete sequence; and large subunit ribosomal RNA gene, partial sequence, with no other matches found in the selected database (All GenBank+EMBL+DDBJ+PDB sequences but no EST, STS, GSS, environmental samples or phase 0, 1 or 2 HTGS sequences). An alignment of partial ssrDNA sequences of several related Microsporidia showed several nucleotide mismatches to that of P. neurophilia present within the 113 base pair amplicon in regions specific to both primers and probe sequences (Figure 3.1). Testing of DNA extracted

from spores of Glugea anomala and Pleistophora hyphessobryconis resulted in no increase in fluorescence detected through 40 cycles of real-time PCR. Sonication Time Study Sonication for 5 min at 55W was determined to be the optimal lysis time to ensure consistently high extraction efficiency of Pseudoloma neurophilia spores with minimal loss of signal due to DNA shearing (Figure 3.2). Extraction Comparison In Trial A, the sonication pretreatment, both unextracted (mean Cq = 26.9) and followed by QIAgen extraction (mean Cq = 27.7), resulted in a lower mean crossing threshold than other methods (Figure 3.3). The multiple comparison with best procedure analysis of Trial B showed chitinase pretreatment followed by QIAgen extraction (CQ), QIAgen extraction with no pretreatment (Q), sonication pretreatment with no further extraction (S) and sonication pretreatment followed by QIAgen extraction (SQ) to be in the best group with sonication alone (S) to have the lowest mean crossing threshold (mean Cq = 28.74). Sonication followed by QIAgen extraction (mean Cq = 29.37) had the next lowest mean crossing threshold (Figure 3.3).

Detection Limit

The assay, using the PN10F/R primer set and probe PN, consistently detected less

than one spore (0.2 spore) per reaction in PBS when the sample was sonicated. This was determined based on the original number of spores (10 spores/100 µl), 2 µl of which were tested by real-time PCR. Pseudoloma neurophilia ssrDNA was detected when spawn water, eggs or sperm samples were spiked with spores at 4-5 spores per real-time PCR reaction in all trials and replicates within trials (Table 3.2) using sonication followed by Qiagen extraction. Parasite ssrDNA was detected in spawn water spiked with as low as 10 spores per liter (0.1 spore per reaction) in one trial, however, it was not detected in the other two. We detected the parasite in eggs spiked with 2 spores per egg (0.4 spore per reaction) in all trials and in all but one replicate. We also detected the parasite in spiked sperm samples, consistently detecting the parasite at 10 spores per µl of sperm (0.5 spore

per reaction). The inconsistent detection at these lower levels of parasite is likely reflective of sampling error inherent in dealing with such dilute concentrations. Comparison to Conventional Assay With the exception of one tissue sample, the real-time PCR assay detected P. neurophilia ssrDNA in at least one log higher dilution compared to the conventional assay (Table 3.3). Analysis of all six samples by the real-time PCR method showed that sonication followed by DNA extraction using the QIAgen DNeasy Blood and Tissue Extraction kit resulted in a mean decrease of crossing threshold of 2.34 compared to the same samples that were not sonicated. Group Spawn Experiment P. neurophilia ssrDNA was detected in the brain and spinal tissues of six of ten adult fish in the group spawning experiment. Eggs were pooled and divided into five aliquots and P. neurophilia ssrDNA was detected in one pool. In contrast, P. neurophilia ssrDNA was detected in nine of ten 1 l water samples from the spawning tank. DISCUSSION Our new real-time PCR test for Pseudoloma neurophilia provides a sensitive test for evaluation of fish tissues and a non-lethal method for detecting spores in fish tissues,

water and in spawning products. This is best illustrated by the decreased crossing

threshold (Cq) of samples using methods employing sonication (Figure 3.3), indicating an increase in the number of ssrDNA copies present. Comparing the test to the conventional PCR based assay developed by Whipps and Kent (2006), we found that it was 10-100 times more sensitive and our calculations indicated that it could detect < 1 spore per reaction based on the number of spores spiked into PBS. This increased sensitivity was not necessarily due to conventional versus real-time format or differences in primers, but more likely due to pretreatment by sonication in our new test as discussed below. Regarding specificity, we designed the primers so that they would be unique to P. neurophilia. Also, we evaluated our test with Glugea anomala from 3-spined sticklebacks, Gasterosteus aculeatus, and Pleistophora hyphessobryconis from neon

tetras, Paracheirodon innesi, as these two Microsporidia might be found in fish research facilities. Three-spined sticklebacks are used in laboratory research and we recently detected P. hyphessobryconis in three zebrafish colonies (Sanders et al., 2010). Additionally, a total of 278 fish from 6 separate populations housed in a known Pseudoloma neurophilia free zebrafish colony were tested using this real-time PCR based assay, with histological analysis on individuals from the same populations performed in parallel (Kent et al., 2011). No fish from this population were positive for P. neurophilia by real-time PCR or histology. Whereas the purpose of the present study was to develop a real-time PCR based assay, we found that the primer set used in this format also will work in a conventional PCR format as a positive/negative screening test. However, we did not determine the sensitivity of the primers using this conventional PCR format. Whereas quantification is not necessary for detection of the parasite, it may be pertinent to implement this feature for future studies on disease progression, transmission and dose response, thus the real- time PCR platform was used. Furthermore, the use of the real-time PCR platform eliminates post-amplification handling of samples, decreasing the chance of cross-

contamination by amplicons leading to false positive results.

Current protocols for the detection of Pseudoloma neurophilia involve either

direct observation of spores in wet mount preparations or routine histological sections using either hematoxylin and eosin or special stains such as Fungi-Fluor (Kent and Bishop-Stewart, 2003), acid fast (Ramsay et al., 2009), or Luna (Peterson et al., 2011). The development of a conventional PCR based test (Whipps and Kent, 2006) has allowed researchers to detect lower levels of the parasite in fish tissues than possible by traditional histological methods and was used effectively to screen brood fish and progeny to establish a specific pathogen free (SPF) zebrafish colony (Kent et al., 2009, 2011). These testing methods are relatively sensitive and specific, but were not evaluated in non-lethal formats. Many populations of zebrafish used in research consist of small numbers of difficult to breed mutant lines of fish and thus it is often impractical to lethally sample statistically significant numbers for diagnostic testing. For example, to screen a

population of 100 fish, assuming a one or greater percent prevalence of infection in the population, 96 fish would need to be examined in order to obtain 95% confidence in detecting at the infection at a prevalence is 1% or greater (Kent et al., 2009; Simon and Schill, 1984). Therefore, detection of P. neurophilia in the fish’s environment or in spawning products offers a practical and desirable alternative. Several studies have been undertaken that incorporated various techniques for concentration and detection of microparasites in water. These include membrane filtration, continuous flow centrifugation and some in combination with immunomagnetic separation or PCR to detect microsporidia and other small parasites in large volumes of drinking or surface water (Bukhari et al., 1998; Fournier et al., 2000; Graczyk et al., 2007; Hallett and Bartholomew, 2006; Hoffman et al., 2007; Swales and Wright, 2000). Graczyk et al (1997) described the use of cellulose-acetate membrane filtration of water followed by dissolution of the filter using acetone to concentrate and visualize Cryptosporidium oocysts. This approach for spore concentration provided a sensitive method for detecting P. neurophilia in water. Successful detection of microsporidia in environmental samples using PCR also requires efficient extraction of DNA from spores. A large portion of our study was,

therefore, devoted to determination of the method that yields the highest amount of PCR

product from spores. Both extraction comparison trials, each utilizing different sample handling, showed that sonication increased the overall sensitivity of the test. In the initial DNA extraction comparison, samples were pretreated in pools and aliquots were taken from these pools for purification in order to minimize the effects of sampling error inherent in dealing with such a small number of spores. We have found it occasionally difficult to obtain homogenous suspensions of P. neurophilia spores and this is likely attributable to some non-polar factors on the exterior of the spore wall, as we often find the spores clustered in wet mount preparations, particularly around the meniscus of air bubbles. The second DNA extraction comparison was based upon the results of the first trial and further confirmed that sonication resulted in much greater DNA extracted even taking into account potential sampling errors. Our results were consistent with those of

Phelps and Goodwin (2007), who showed that the amount of DNA obtained from another egg-associated fish microsporidium, Ovipleistophora ovariae, was increased over 500 times by sonicating spores compared to proteinase K digestion alone. As with their study, we observed numerous intact spores in preparations following the QIAgen extraction, indicating that this method and others which depend solely on proteinase K and detergent digestion is not effective for disrupting some microsporidian spores. This is further supported by the higher overall sensitivity obtained from tissue samples which had been sonicated prior to DNA extraction. Although the extraction comparison showed that sonication without the additional step of DNA extraction using the QIAgen kit had the highest sensitivity with purified spores, this step was necessary for extraction from tank water and eggs due to inhibitory elements found in these sample matrices as determined by spiking sonicated, unpurified samples (data not shown). Whereas this inhibition could potentially be eliminated by diluting the samples or adding adjuncts such as bovine serum albumin, the DNA extraction step was added in order to maintain a consistent extraction with samples potentially containing variable amounts of inhibitory substances.

Our test was also effective for spawning water, eggs and sperm. We were able to

detect very low levels of the parasite in replicates within trials and obtained similar

results with independent trials. Zebrafish are spawned by placing pairs or groups into small tanks with screen bottoms overnight. Using the F1 progeny of surface-disinfected eggs is the main source for establishing new populations of zebrafish and introducing new lines into existing zebrafish colonies (Lawrence, 2007), an approach that has been used for decades to avoid movement of salmonid pathogens (Kent and Kieser, 2003; Stead and Laird, 2002). For the latter, sex products are also screened for specific pathogens that are maternally transmitted (Miriam et al., 1997). The use of surface disinfected eggs is likely to be effective for avoiding certain bacterial pathogens but this practice has not stopped the spread of Pseudoloma neurophilia within zebrafish colonies. The reason for this is likely twofold; the ineffectiveness of levels of chlorine used for disinfection of eggs (Ferguson et al., 2007) and the maternal transmission route of the

parasite. Other fish microsporidia are vertically transmitted (Phelps and Goodwin, 2008), and several lines of indirect and observational data indicate that there is a risk of maternal transmission with P. neurophilia, either transovum (within eggs) or transovarial (outside egg at spawning): 1) the parasite has been observed in several facilities that have been established and only use F1 progeny from disinfected eggs, 2) the parasite is common in the ovaries and occasionally within eggs, 3) larval zebrafish are highly susceptible to the infections. We detected the parasite in water and eggs from a spawn tank of a population of brood fish with an infection prevalence of 60%, providing further evidence of maternal transmission. Given that resistant spores are abundant at spawning, the only way to reliably avoid maternal transmission is to identify infected broodfish or progeny with a highly sensitive diagnostic test. Screening of 10 day old fry and broodstock from several zebrafish lines were used to establish a P. neurophilia SPF facility at Oregon State University (Kent et al., 2009, 2011). Our test also has potential to be used for nonlethal screening of adult stocks. This non-lethal format is feasible as adult zebrafish spawn prodigiously and paired spawning is often performed in 1 l or less of water (Harper and

Lawrence, 2010; Lawrence, 2007). The results of the group spawn experiment illustrate

the potential application of this assay in this format. However, it is important to note that

testing sex products and water may not be reliable for detecting the parasite in all infected fish. The parasite does not occur in the ovaries of all infected female zebrafish (Kent and Bishop-Stewart, 2003) and thus these fish may be less likely to shed spores at spawning. Also, we have not seen the parasite in the testis by histology, and thus this approach for non-lethal testing may not be useful for identifying infected males. With these limitations, the test would still be of value in that a positive result from pooled samples from a group spawn would clearly demonstrate that at least one fish in the spawning group was positive. Moreover, our test is extremely sensitive, and thus a negative result with spawning water or eggs would be highly suggestive that the progeny from this particular spawn were not infected. Other experiments performed in our laboratory have produced similar results and further studies are currently underway to

elucidate the progression and transmission of the parasite within populations of fish in order to determine the predictive value of this type of test and its reliability in detecting positive fish in lightly infected populations. We are also further investigating the role of males in maternal transmission of the parasite. Cryopreserved sperm is used for long term storage of zebrafish lines and is frequently used to establish new populations (Westerfield, 2007). Whereas we have not detected the parasite in testis by histology, sperm could be contaminated during the squeezing process to obtain sperm. Hence, our new test provides a method for screening sperm for the parasite. In conclusion, we provide here a new real-time PCR based assay for P. neurophilia that is more sensitive that our previous test. Additionally, we have developed a method by which to sample and detect the parasite in water, eggs, and sperm, thus providing the foundation for a non-lethal test. It is intended that personnel involved in the maintenance of laboratory Danio rerio populations use this protocol as a basis for their own testing protocols and modify it to suit the needs of individual facilities for monitoring and screening. ACKNOWLEDGEMENTS

This study was supported by grants from the National Institutes of Health (NIH

NCRR 5R24RR017386-02 and NIH NCRR P40 RR12546-03S1). We thank Dr. C.

Whipps, State University of New York, and Dr. T. Peterson, Oregon State University, for manuscript review, C. Buchner of the Sinnhuber Aquatic Research Laboratory for providing negative control specimens, and G. Weaver, Oregon State University Department of Statistics, for statistical support.

Fig. 3.1. Partial ssrDNA sequence alignment of Pseudoloma neurophilia and related microsporidia. Pseudoloma neurophilia (Genbank AF322654), Loma embiotocia (Genbank AF320310) , Ovipleistophora mirandellae (Genbank AF356223), Loma salmonae (Genbank U78736), Glugea sp. (Genbank AY090038), Loma morhua (Genbank GQ121037), Loma psittaca (Genbank FJ843104), Pleistophora hyphessobryconis (Genbank GU126672), Heterosporis anguillarum (Genbank AF387331), Ichthyosporidium giganteum (Genbank L13430), Pleistophora mulleri (Genbank FN434084), Dasyatispora levantinae (Genbank GU183263), Glugea stephani (Genbank AF056015), Loma salmonae (Genbank HM626215), Loma sp. (Genbank HM626217), Glugea anomala (Genbank AF044391), Loma sp.(Genbank AF104081), Glugea atherinae (Genbank U15987), Loma acerinae (Genbank AJ252951), Pleistophora typicalis (Genbank AJ252956), Pleistophora sp (PA) (Genbank AJ252958), and Ichthyosporidium sp.(Ganbank L39110) Pn10F and Pn10R primer locations are underlined and PN10 probe location is shaded. Asterisks denote regions of nucleotide similarity.

Fig. 3.1. Partial ssrDNA sequence alignment of Pseudoloma neurophilia and related microsporidia. (Continued)

22 24

26 28 30 32 34 36 Quantification Cycle (Cq) 38 40 0 15 30 60 300 600 900 Sonication Time (seconds)

Fig. 3.2. Pseudoloma neurophilia spore sonication time study. Quantification cycle thresholds (Cq) of triplicate suspensions of 1000 spores of P. neurophilia sonicated at

55W for different time points and tested by real-time PCR. Bars represent mean Cq;

Points represent individual sample Cqs.

Fig. 3.3. Pseudoloma neurophilia spore extraction method comparison by quantification cycle (Cq). NT: no treatment, M: MoBio, Q: QIAgen, P: peroxide, P+M: peroxide + Mobio, P+Q: peroxide + QIAgen, S: sonication, S+M: sonication + Mobio, S+Q: sonication + QIAgen, B+Q: bead beating + QIAgen, C+Q: chitinase + QIAgen. Bars represent mean Cq; points represent actual Cq for individual samples. Sonication treatment alone showed the lowest mean Cq in Trial A, 26.74 followed by sonication with DNA extraction by QIAgen silica gel membrane method (mean Cq = 27.73). Trial B results showed the lowest mean Cq for sonication alone, 28.74, followed by sonication

and QIAgen DNA extraction combined, 29.93.

Table 3.1. Experimental setup for two extraction comparison trials. Trial A samples represent 300 µl aliquots taken from an initial pool of 30,000 spores in 900 µl of PBS. Following pretreatment, samples were divided into 6 aliquots for DNA extraction. Each extraction method was performed twice. Samples in Trial B represent individual purified spore suspensions containing 1,000 spores each in 100 µl PBS. Methods in Trial B were performed in triplicate.

DNA Trial Sample Pretreatment Extraction

A 1 NT No Extraction

QIAgen

MoBio

2 Hydrogen No Extraction Peroxide QIAgen MoBio

3 Sonication No Extraction QIAgen

MoBio

B 1 None None 2 Bead Beating QIAgen 3 Chitinase QIAgen 4 None QIAgen 5 None MoBio 6 Sonication None 7 Sonication QIAgen

Table 3.2. Pseudoloma neurophilia spiked sample results from three separate trials. Numbers for each trial represent quantification cycle threshold (Cq) detected by real-time PCR per sample. Spawn water consists of purified spores of P. neurophilia in 1 l of water from a group spawn of P. neurophilia free (SPF) zebrafish. Eggs consist of purified spores of P. neurophilia in a 40 µl aliquot (representing approximately 20 eggs based on an initial volume of 500 eggs/1ml) of homogenate made from eggs spawned by SPF zebrafish. Sperm samples consist of purified spores of P. neurophilia in a 5 µl aliquot of sperm obtained by squeezing SPF zebrafish males. - : no copies of P. neurophilia ssrDNA detected after 40 cycles.

Starting Mean Sample Concentration Trial 1 Trial 2 Trial 3 (Standard Type (Spores per Deviation) reaction)

Spawn

Water 500 ( 5 ) 36.7, 35.0, 36.4 35.8, 34.8, 37.9 35.6, 36.1, 37.9 36.2(1.1) 100 ( 1 ) -,-,39.9 39.4, 37.4, 38.8 38.8,39.3,39.7 39.0(0.8) 50 ( 0.5 ) 38.3, 37.5, 39.4 38.9, -, 37.3, -, 39.4 38.5(0.9)

10 ( 0.1 ) -,-, -,-, 38.1, 38.5, 38.3(0.3)

Eggs 400 ( 4 ) 35.6, 36.3, 35.3 36.6, 37.9,36.7 36.1, 36.5,36.2 36.4(0.7) 40 ( 0.4 ) 38.6, -, 36.2, 36.8, 36.0 35.2, 36.9, 36.9 36.7(1.1)

4 ( 0.04 ) -,-, 37.2, 38.6, 37.3 36.6, 37.6, 37.5(0.7)

Sperm 500 ( 5 ) 32.7, 32.3, 31.9 35.5, -, 34.4 32.8,32.1, 31.3 32.9(1.4)

50 ( 0.5 ) 35.9, 36.6, 34.8 38.0, -, 34.4 35.6, 35.6, 34.9 35.7(1.2)

5 ( 0.05 ) 36.4, -, 36.3 36.8, -, 34.4 35.1,-, 35.8(1.0)

Table 3.3. Comparison of Pseudoloma neurophilia detection by new real-time PCR assay utilizing sonication pretreatment of samples with conventional PCR assay described by Whipps and Kent (2006). The real-time PCR assay described detected the parasite from one to three log dilutions higher than the conventional assay (+ = P. neurophilia ssrDNA detected, - = No P. neurophilia ssrDNA detected).

Individual Conventional PCR Real-time PCR 0 -1 -2 -3 -4 0 -1 -2 -3 -4 1 + - - - + + + - 2 + - - - + - - - 3 + + - - + + + + 4 + - - - + + + - 5 + - - - + + - - 6 + - - - + + - -

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Hallett, S.L., and Bartholomew, J.L. (2006). Application of a real-time PCR assay to detect and quantify the myxozoan parasite Ceratomyxa shasta in river water samples. Dis. Aquat. Organ. 71, 109.

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Phelps, N.B.D., and Goodwin, A.E. (2008). Vertical transmission of Ovipleistophora ovariae (Microspora) within the eggs of the golden shiner. J. Aquat. Anim. Heal. 20, 45– 53.

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Williams, B.A.P., Lee, R.C.H., Becnel, J.J., Weiss, L.M., Fast, N.M., and Keeling, P.J. (2008). Genome sequence surveys of Brachiola algerae and Edhazardia aedis reveal microsporidia with low gene densities. BMC Genomics. 9, 200.

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Chapter 4

Verification of intraovum transmission of a microsporidium of vertebrates: Pseudoloma neurophilia infecting the zebrafish, Danio rerio

Justin L. Sanders1*, Virginia Watral1, Keri Clarkson2, Michael L. Kent1,2

PLOS One, In press San Fancisco, CA, USA

1Department of Microbiology, Oregon State University, Corvallis, OR 97331, USA 2Department of Biomedical Sciences, Oregon State University, Corvallis, OR 97331, USA

ABSTRACT Direct transmission from parents to offspring, referred to as vertical transmission, occurs within essentially all major groups of pathogens. Several microsporidia (Phylum Microsporidia) that infect arthropods employ this mode of transmission, and various lines of evidence have suggested this might occur with certain fish microsporidia. The microsporidium, Pseudoloma neurophilia, is a common pathogen of the laboratory zebrafish, Danio rerio. We previously verified that this parasite is easily transmitted horizontally, but previous studies also indicated that maternal transmission occurs. We report here direct observation of Pseudoloma neurophilia in the progeny of infected zebrafish that were reared in isolation, including microscopic visualization of the parasite in all major stages of development. Histological examination of larval fish reared in isolation from a group spawn showed microsporidian spores in the resorbing yolk sac of a fish. Infections were also observed in three of 36 juvenile fish. Eggs from a second group spawn of 30 infected fish were examined using a stereomicroscope and the infection was observed from 4 to 48 hours post-fertilization in two embryos. Intraovum infections were detected in embryos from 4 of 27 pairs of infected fish that were spawned based on qPCR detection of P. neurophilia DNA. The prevalence of intraovum infections from the four

spawns containing infected embryos was low (~1%) based on calculation of prevalence

using a maximum likelihood analysis for pooled samples. Parasite DNA was detected in the water following spawning of 11 of the infected pairs, suggesting there was also potential for extraovum transmission in these spawning events. Our study represents the first direct observation of vertical transmission within a developing embryo of a microsporidian parasite in a vertebrate. The low prevalence of vertical transmission in embryos is consistent with observations with some other fish pathogens that are also readily transmitted by both vertical and horizontal routes.

INTRODUCTION Pathogens employ a range of different mechanisms to infect new hosts. Vertical or maternal transmission is characterized by pathogens being transmitted to progeny from parents, usually through an infected female host. This mode of transmission is employed by some obligate parasites, which cannot complete their life cycles without a host. The methods by which pathogens are transmitted can exert a powerful influence on virtually all aspects of the biology of the host, either directly or indirectly. Indeed, it was argued that pathogens and their effects on hosts are responsible for nearly all aspects of biological organization (Herre, 1993). The population structure of the host species has an important influence on mode of transmission employed by pathogens (Ebert and Herre, 1996; Herre, 1993). For example, in populations where individuals are spatially separated, minimizing the opportunity for horizontal transfer, selection would favor maternal, or vertical, transmission, ensuring propagation of the parasite. The mode of transmission also has a direct influence on the virulence of a parasite with selection favoring levels of parasite reproduction, and thus virulence, that provide the highest level of fitness for the parasite. Pathogens which are transmitted vertically

are generally less virulent that those which are transmitted horizontally as there is

selective pressure for the survival of the infected female to live to reproductive age and

pass the infection on to progeny (Ewald, 1987). As a result, theses parasites generally do not proliferate to high numbers in the host, minimizing disease and thus host mortality. The virulence-related characteristics of vertically transmitted pathogens are generally limited to those that increase the number of susceptible hosts, including feminization of male hosts (Dunn et al., 2001). Microsporidia are obligate intracellular pathogens with species infecting virtually all animal phyla. This distinctive group has undergone numerous taxomonic revisions since first being described (Corradi and Keeling, 2009). Originally assigned to the Schizomycetes, a group containing yeast-like fungi (Nageli, 1857), the Microsporidia were subsequently moved to other groups such as the Sporozoa (Balbiani, 1882) and Archaezoa (Cavalier-Smith, 1983). Their assignment to the Archaezoa, a group of

protists considered to be “primitive” due to the absence of some typical eukaryotic features, was based mainly on their apparent lack of mitochondria and was further supported by early molecular phylogenetic analyses based on small subunit ribosomal RNA gene sequences. This was, however, later found to be in error as subsequent molecular and ultrastructural analyses have shown the presence of relictual mitochondria (Williams et al., 2002) and more sophisticated phylogenetic analyses that account for rate variation and the presence of numerous fungal-type products, notably trehalose and chitin, placed the Microsporidia once again among the Fungi. Their exact placement among the Fungi (i.e., as early-branching or sister to the Fungi) is currently debated (Williams and Keeling, 2011). Microsporidia have been shown to employ a diverse range of transmission strategies. Several microsporidian species are transmitted horizontally, generally by ingestion of spores from either water contaminated by feces or tissue from dead infected hosts (Canning et al., 1986). Maternal transmission of microsporidian parasites has been noted for several species infecting crustaceans (Terry et al., 2004) and insects (Becnel and Andreadis, 1999). Whereas some microsporidian species are only vertically transmitted (Terry et al., 1999), many are transmitted both horizontally and vertically.

Maternal transmission occurs by two general mechanisms, which in arthropods are

characterized as transovarial (within the egg) and transovum (parasite is shed outside of the egg during egg laying/spawning). These terms were developed to describe vertically transmitted microsporidia of insects, and are somewhat confusing when applied to vertebrates. Hence, we refer to these two types of transmission as intraovum and extraovum, respectively. Regarding vertebrates, the earliest recorded evidence of vertical transmission comes from Hunt et al (Hunt et al., 1972) who observed Encephalitozoon cuniculi infections in gnotobiotic rabbits, strongly suggesting transplacental transmission of this parasite. Also, several lines of indirect evidence support vertical transmission of microsporidia in fishes. The strongest evidence of vertical transmission in fish comes from Ovipleistophora ovariae, a microsporidium that infects the golden shiner,

Notemigonus crysoleucas, and has been observed to infect only female fish. In this species, spores are found almost exclusively within the ovaries and developing oocytes (Summerfelt, 1964). While large amounts of O. ovariae DNA has been detected within surface-decontaminated, spawned eggs and developing larval fish (Phelps and Goodwin, 2008), the parasite has not been directly observed in the progeny of infected fish. Ovipleistophora mirandellae also infects the oocytes and eggs of cyprinid fishes (Vaney and Conte, 1901) and thus has been suggested (Maurand et al., 1988) to undergo intraovum transmission. Loma salmonae is a microsporidian pathogen of salmon and trout, and infects the ovigerous stroma, but not eggs (Docker et al., 1997). The translocation of Loma salmonae to fish farms in Chile which had been populated using only surface decontaminated, fertilized eggs is also suggestive of vertical transmission (Brown et al., 2010; Docker et al., 1997). The zebrafish, Danio rerio, is an important laboratory model for toxicology, developmental biology, cancer, and infectious disease research. It has a well- characterized immune system (Traver et al., 2003) and the complete genome has been sequenced with the genetic map showing an overall highly conserved synteny with the human genome (Postlethwait et al., 1998). A microsporidium, Pseudoloma neurophilia,

is responsible for chronic infections of zebrafish (Matthews et al., 2001; Sanders et al.,

2012) and is common in zebrafish research colonies (Murray et al., 2011). The high prevalence of a microsporidium in laboratory zebrafish populations provides an ideal opportunity to learn more about the transmission characteristics of this parasite. As the name implies, Pseudoloma neurophilia chronically infects neural tissue and develops into mature spores mainly in the hindbrain, spinal cord, motor nerve ganglia and spinal nerve roots. The parasite enters the host via the intestinal epithelium and infects extraintestinal skeletal muscle myocytes in early stages of the infection (Cali et al., 2012). Infections by the parasite result in a spectrum of disease ranging from minimal to no clinical presentation to acute mortality. We have shown that P. neurophilia is horizontally transmissible by bath exposure and cohabitation with live fish, ostensibly per os. There is growing evidence of vertical transmission for this parasite as follows: 1. The presence

of microsporidian spores visualized by histology in the ovaries and developing oocytes of infected adult female fish (Kent and Bishop-Stewart, 2003), 2. P. neurophilia DNA detected by PCR in spawn water and eggs from infected adults (Sanders and Kent, 2011), 3. High numbers of P. neurophilia spores in ovaries of infected females (Sanders et al., 2012), 4. Repeated detection of P. neurophilia in the progeny of two lines of zebrafish screened as part of a protocol to develop a specific pathogen free colony leading to the exclusion of these lines from the colony (Kent et al., 2011). Despite the significant indirect evidence, observation of the intra-ovum transmission of a microsporidian parasite in a vertebrate has not been verified. By utilizing a combination of a qPCR method (Sanders and Kent, 2011) and microscopy on a number of spawning fish and progeny, we provide direct evidence of the intraovum transmission of P. neurophilia. We also provide additional data supporting the hypothesis that extraovum maternal transmission is also an important route of transmission. RESULTS Group spawns

One of 24 egg pools was positive for P. neurophilia DNA. Histological

examination of 7 dpf larval fish revealed the presence of microsporidian spores in the

epidermis (Figure 4.1A) and resorbing yolk sac (Figure 4.1B) of one of 112 fish examined. Histological examination of juveniles from this spawn, reared in isolation, at 8 wk post fertilization revealed spores in various tissues in 3 of 36 fish examined, such as the lamina propria of the intestine (Figure 4.1C) and the ovigerous stroma (Figure 4.1D), and inner ear. Eggs from a second group spawn of 30 adults from a population of infected fish were examined using a stereomicroscope with a transmitted light source. Distinctive opaque regions were seen in two embryos at approximately 4 hours post-fertilization (Figure 4.2A). One embryo was maintained at 28°C and examined again at 24 and 48 hours post fertilization during which time it appeared to develop normally (Figures 4.2B, 4.2C). The other embryo was sacrificed and examined by wet mount microscopy. Upon

examination using higher magnification, these regions were found to consist of large aggregates of refractile spores with characteristic polar vacuoles, consistent in shape and size to Pseudoloma neurophilia (Figure 4.2D). The embryo was then homogenized in sterile water and the spores quantified by hemocytometer. Thirty thousand spores were present in this particular embryo. Prevalence of P. neurophilia within oocytes and detection in spawning water Pseudoloma neurophilia DNA was detected within eggs from 4, and in the spawn water of 11, of 27 paired spawns (Table 4.1). The mean prevalence of P. neurophilia within spawned eggs from these 4 positive spawns was calculated to be 0.9% (CI 0.06 – 3.45%). Histological examination of spawning pairs Spores were observed in the spinal cords and brains of 43 out of 54 spawning adult fish that were examined. The parasite was observed in the ovigerous stroma of 7 of 27 females, and within developing follicles of 4 of these 27 fish (Table 4.1). Of the 4 females with eggs that tested positive by qPCR after spawning, spores were observed in the ovigerous stroma of two and within a developing oocyte of one. No spores were

observed in the testes of any male fish examined nor in any tissues of fish in which

spores were not also observed in the spinal cord.

Dose response of larval fish to P. neurophilia Larval fish in two trials became infected when exposed to 300 or 500 spores/ml, but not at lower concentrations (Table 4.2). Whereas there was variability between replicates and trials, owing primarily to the high numbers of mortalities observed during the duration of the experiment, there was a trend for a dose response. DISCUSSION A combination of indirect (qPCR) and direct (microscopy) methods demonstrated intraovum transmission of the microsporidian parasite, Pseudoloma neurophilia, in its zebrafish host. Evidence for the vertical transmission of fish pathogens has often been indirect or circumstantial, especially for those that can be transmitted both horizontally and vertically. Determination of whether vertical transmission occurs, and by which

mechanism (i.e., intraovum versus extraovum) can be complicated by several factors including low overall prevalence of vertical transmission, variation of occurrence both within and between clutches of eggs, and the ability of many of these pathogens to survive for long periods in the water leading to extraovum transmission. Here we verify for the first time intraovum transmission of microsporidium infecting a vertebrate host. There are several similarities and some differences between P. neurophilia and Ovipleistophora ovariae. The latter microsporidium was first described in ovarian infections of the golden shiner, Notemigonus crysoleucas, another cyprinid fish that is an important bait fish raised in aquaculture in the United States. It is found with high prevalence in females from commercial fish farms and has been shown to greatly reduce fecundity as the fish ages. Ovipleistophora ovariae is found primarily in the ovarian stroma and within intermediate to fully mature oocytes. However, it has also been observed in liver and kidney tissue from a few infected females (Summerfelt and Warner, 1970). It has not been observed in tissue from male fish, however DNA from the parasite has been detected in testes (Phelps and Goodwin, 2007). Horizontal transmission of O. ovariae has been demonstrated by feeding fry and fingerlings spores adsorbed to feed

(Summerfelt, 1972). By treating eggs from infected shiners with RNase Away, as done

in our study, Phelps and Goodwin (2008) detected high numbers of copies of O.

ovariae DNA in all pools of eggs tested, providing strong evidence of intraovum transmission of the parasite. Fry hatched from the same clutch of eggs also showed high levels of O. ovariae DNA at 48 hours post hatch. However, no parasites were observed in these fish by microscopy. The parasite almost completely replaces the egg interior with most infected eggs, and thus Summerfelt (1972) suggested that embryos from these eggs are not viable, but rather serve as a source of infection to siblings. In this case, therefore, O. ovariae is actually transmitted by extraovum maternal transmission because the infected egg itself does not directly result in an infected fish. In contrast to O. ovariae, relatively few zebrafish eggs and embryos are infected by P. neurophilia. We were able to observe spores of Pseudoloma neurophilia in one 7 day post fertilization fish (i.e. 5 day post-hatch) by histology, and this low prevalence

correlates with that found by screening eggs by qPCR (~1%). While we are unable to differentiate between merogonic or spore stages of P. neurophilia using the qPCR assay, the observation of mature spores within a developing embryo and developing oocytes confirms the presence of this stage in at least some cases of intraovum transmission. This does not preclude the presence of presporogonic stages within oocytes, which are more difficult to visualize by standard histological methods. The other member of the genus Ovipleistophora, O. mirandellae, also infects ovaries, testis, and eggs of several fishes (Maurand et al., 1988; Pekkarinen et al., 2002). It has been suggested to be vertically transmitted, but we are not aware of empirical data supporting this hypothesis. There are other examples of fish pathogens that, like P. neurophilia, are transmitted both horizontally and vertically, and show a low prevalence of infected eggs. It is not necessary to have a high prevalence of infected eggs and embryos within a clutch to infect the next generation as vertical transmission is followed by robust horizontal transmission within the F1 siblings. Two bacterial and one viral pathogen of salmonid fishes use these modes of transmission. Renibacterium salmoninarum, for example, is an obligate Gram positive bacterial pathogen that has been spread around the world with egg

shipments (Fryer and Sanders, 1981). It was present within surface disinfected eggs from

experimentally infected rainbow trout Onchorynchus mykiss at a prevalence of 1.7%

(Bruno and Munro, 1986), and from two heavily infected coho salmon Oncorhynchus kisutch from separate studies at prevalences of 5.8 % (Evelyn, 1984) and 11.6% (Evelyn et al., 1984). Similarly, 13% of newly spawned eggs from infected steelhead trout, Onchorynchus mykiss, were found to be infected with Flavobacterium psychrophilum (Brown et al., 1997), a serious pathogen in salmon and trout hatcheries (Holt et al., 1993). The small size of these bacteria allow for their passage into the egg after it is released from the female through the egg micropyle prior to fertilization, whereas the larger size of microsporidian spores such as P. neurophilia (approximately 3.5 µm by 5 µm) prevents it from entering through the approximately 1.7 – 2.5 µm diameter micropyle of the developed zebrafish egg (Hart and Donovan, 1983), requiring it to be present in the

oocyte prior to spawning. This could account for the somewhat higher prevalence of intraovum transmission observed with these bacteria. Infectious hematopoietic necrosis virus (IHNV), a virus infecting salmonids, was responsible for the failure of successful culture of sockeye salmon, Oncorhynchus nerka, in Alaska prior to 1981 (Meyers and Thomas, 1990). The implementation of a risk management approach consisting of the use of virus-free water supplies, rigorous disinfection, and compartmentalization of eggs and fry to contain virus outbreaks when they occur was unsuccessful in completely resolving IHNV epizootics (Meyers, 1998). One possibility for this failure was suggested to be that intraovum vertical transmission of the virus occurs, resulting in a few subclinically infected progeny. There has been evidence to support the intraovum transmission of IHNV. Mulcahy and Pascho (1985) described the isolation of IHN virus from fry and eggs held in the laboratory under virus- free conditions in two occurrences. They found that only a small proportion of eggs and fry tested contained IHN from one female with a high titer present in the body cavity. The low prevalence of intraovum transmission of P. neurophilia observed is adequate to maintain the infection between generations, given the large numbers of larval

siblings in a single spawn and the high rates of horizontal transmission. Indeed,

observing the infection by histology, we detected P. neurophilia in only one 7 dpf larva,

but within 8 weeks increased to 11%, including spores developing within the ovigerous stroma of one fish. Following initial infection, the parasite employs several routes of transmission (Sanders et al., 2012). Waterborne transmission occurs by co-habitation with infected fish and by ingestion of infected tissue. Zebrafish spawn every few weeks, releasing spores and infected eggs into the water which are then fed upon by tanks mates. Also, feeding on infected carcasses is an efficient route of transmission. Zebrafish do not generally attack other fish in an aquarium or school, but quickly scavenge dead fish. In addition to intraovum transmission, Pseudoloma neurophilia, as well as IHN and Renibacterium salmoninarum, employs a second maternal transmission mechanism; extraovum transmission. High levels of P. neurophilia in the ovaries and in spawn water have been reported previously (Sanders and Kent, 2011; Sanders et al., 2012), and with

our paired spawn analysis the spawn water contained the parasite more often than eggs. Also, the dose response experiment agreed with the previously studies (Ferguson et al., 2007) showing that larvae, which begin feeding about 4-5 d after fertilization, are very susceptible. Again this is similar to Renibacterium salmoninarum, which is often found at high prevalence and concentration in the ovarian fluid from infected female salmon (Pascho et al., 1998). Whereas very few eggs were internally infected by this bacterium, it was detected on the surface of 38 % of the eggs (Bruno and Munro, 1986). This strategy of extraovum transmission likely occurs with the IHN virus as well, and extends to infecting the next generation beyond the infected parents own progeny in a given river or stream (LaPatra et al., 1991; Traxler et al., 1997). Traxler et al. (1997) showed no correlation between infection levels in ovarian fluid compared to eggs, indicating that extraovum transmission is the major route of vertical transmission for this viral pathogen. The mode of vertical transmission we report has practical implications. Extraovum transmission in fish can be mitigated by the use of rigorous surface decontamination of eggs whereas intraovum transmission (i.e., pathogens present within the egg) would render any surface decontamination method ineffective. This approach has been used to avoid extraovum viruses and bacteria following spawning with

salmonids (Kent and Kieser, 2003; Stead and Laird, 2002), but is ineffective for P.

neurophilia (Ferguson et al., 2007). The high prevalence of P. neurophilia in laboratory zebrafish colonies, the risk of intraovum transmission, and the lack of disinfectants that will kill spores but not eggs (Ferguson et al., 2007) led to the development of a zebrafish colony which is specific pathogen free (SPF) for P. neurophilia (Kent et al., 2011), accomplished by rigorous screening of brood fish and progeny for the pathogen with our PCR tests (Sanders and Kent, 2011; Whipps and Kent, 2006). The population of P. neurophilia used in this study has been maintained for > 4 years by horizontal passaging in zebrafish held in aquaria (i.e., exposing fish to infected spinal material, cohabitation with infected fish). This may have resulted in the selection of parasite strains which are most efficiently transmitted horizontally, resulting in the very low prevalence of intraovum transmission observed in the current study. Intraovum

transmission does occur nonetheless, and the occurrence of this method of transmission could likely be experimentally manipulated. Several species of Microsporidia are transmitted both horizontally and vertically (Dunn et al., 2001; Smith and Dunn, 1991). Even among species which are transmitted solely transovarially, transmission efficiency has been observed to be variable between broods (Smith and Dunn, 1991). The zebrafish has been used to study host-pathogen interactions of numerous bacterial and viral pathogens (Phelps and Neely, 2005; van der Sar et al., 2004; Sullivan and Kim, 2008). Pseudoloma neurophilia infections of zebrafish would provide a good model to investigate the evolution of vertical transmission as it employs both horizontal and vertical transmission. We summarize the transmission of P. neurophilia in Figure 4.3: The primary mode of transmission appears to be horizontal, with many numbers of P. neurophilia spores being released into the water from ovaries of females during spawning (Fig. 4.3a), which occurs on a routine basis every few weeks. Feces and urine are also a source of infection from live fish. This is supported by the presence of P. neurophilia spores developing in the renal tubules of (Fig. 4.3b). Zebrafish are relatively docile fish, but quickly scavenge dead tank mates, providing another route of horizontal transmission.

Alternatively, vertical transmission, either intraovum (Fig. 4.3c) or extraovum

(Fig. 4.3d) can occur via the female host. Spores shed by females in the ovarian fluid during spawning can then be ingested by the developing larvae, leading to new infections. Intraovum transmission can lead to death of the infected embryos (Fig. 4.3e) or directly to viable, infected hosts (Fig. 4.3f) which can subsequently infect other hosts by horizontal transmission and later, by vertical transmission. Whereas the occurrence of intraovum transmission appears to be rare, it is nonetheless important for maintenance of the parasite in laboratory zebrafish colonies especially considering that infected embryos appear to be able to develop normally, at least to the post-hatch larval stage.

METHODS All experimental protocols for the procedures using zebrafish were approved by the Oregon State University Institutional Animal Care and Use Committee (Proposal Number: 4148). Dose response in larval fish. Larval zebrafish have previously been shown to be highly susceptible to Pseudoloma neurophilia, with an estimated 310 spores/fish exposure resulting in 2 of 6 fish becoming infected at 7 days post exposure (Ferguson et al., 2007). In order to further assess the potential for extraovum transmission of P. neurophilia from spawned ovarian fluid to progeny, 5 dpf larval zebrafish were exposed to low numbers of the parasite and examined by wet mount for the presence of microsporidian spores developing within tissues. Two separate dose-response trials were conducted. Zebrafish embryos (AB line) were obtained from the P. neurophilia specific pathogen free zebrafish colony housed at the Sinnhuber Aquatic Research Laboratory at Oregon State University (Kent et al., 2011). Embryos were placed in sterilized 100 ml glass beakers (a total of 6 beakers) in groups of 25 embryos/beaker and held in 25 ml of autoclaved

system water at 28°C. At 4 dpf, larval fish were fed 500 µl of a concentrated

paramecium suspension and four groups were inoculated with a suspension of

P.neurophilia spores obtained from infected adult fish at a concentration of 300 spores/ml. At 7 days post exposure, all fish were euthanized by an overdose of Finquel (MS-222, Argent Laboratories, Redmond, WA, USA) and examined microscopically by wet mount. The presence of spores was recorded for individual fish. Infections in progeny from group spawns Whereas the microsporidium has been observed in eggs at various stages of development and detected by PCR in eggs after spawning (Sanders and Kent, 2011), it has not been visualized in embryos prior to hatching. Therefore, to obtain large numbers of potentially infected eggs, we harvested eggs that were obtained using group spawning from a known population of infected fish.

One population of presumed infected fish was spawned (approximately 75 fish) in a 10 L tank with a nylon mesh false bottom insert. Fish were held at 28°C overnight. Eggs were collected (approximately 2000), rinsed with sterile system water, placed in 90 mm glass petri dishes (approximately 200 eggs/dish) and held at 28°C in sterile embryo media (Westerfield, 2007). At 2.5 dpf, 24 pools of ten eggs each were collected and placed in a 1.5 ml microcentrifuge tube. DNA was extracted from the egg pools and then tested by qPCR using the method described in Sanders & Kent (2011). At 7 dpf, 112 fish were euthanized by an overdose of Finquel and fixed overnight in Dietrich’s fixative. Fish were then placed in a 4 X 6 agarose array (Sabaliauskas et al., 2006), topped with molten agarose and processed for histology. Fifteen serial 5 µm sections were cut from each array, stained with the Luna stain (Peterson et al., 2011), and examined by light microscopy. At 8 weeks old, the remaining fish were euthanized and fixed in Dietrich’s fixative. Thirty-six fish were processed for histology, stained with the Luna stain and examined as described above. A second population of 30 infected adult fish was spawned. Eggs were examined for the presence of opaque regions potentially indicative of aggregates of microsporidian spores using a stereomicroscope with a transmitted light source.

Infections in eggs and spawn water from fish spawned in pairs

In order to determine the prevalence of P. neurophilia present within eggs after spawning, we applied the method described by Phelps and Goodwin (2008) to determine the prevalence of Ovipleistophora ovariae within golden shiner eggs. Adult zebrafish that were previously exposed to P. neurophilia by feeding infected material were spawned in pairs in 1 L of water in a tank with a false-bottom insert. Successfully spawning pairs were euthanized by an overdose of Finquel and fixed in Dietrich’s fixative for subsequent processing and histological examination. Single sagittal sections were cut from these adult fish, stained with the Luna stain, and examined for the presence of P. neurophilia spores. Water from the spawning tanks was collected and filtered through a 1.2 µm nitrocellulose filter and processed as previously described (Sanders and Kent, 2011). Eggs were collected and placed in pools ranging from 10-30 eggs,

depending on the total number of eggs collected from each clutch, in 1.5 ml tubes. The commercial DNA/RNA decontaminating product RNase AWAY (Molecular BioProducts, Inc., San Diego, California, USA) was added at full concentration to pools of eggs in order to destroy any P. neurophilia spores or DNA present on the exterior of the eggs. After 10 minutes, the RNase AWAY was aspirated, the eggs were rinsed twice with DNA-free water and DNA was extracted from the egg pools as previously described (Sanders and Kent, 2011) with the exception that sonication was performed in a Bioruptor sonicating bath (Diagenode, Denville, NJ, USA) at high power for 14 minutes (30s on, 30s off at 4°C) prior to digestion with lysis buffer. Phelps and Goodwin (2008) determined that the RNase AWAY treatment removed DNA of Ovipleistophora ovariae spores outside the eggs of golden shiners. To verify that this method is efficacious for P. neurophilia spores outside the eggs of zebrafish, we conducted the following. Eggs were obtained from a population of P. neurophilia-free zebrafish and placed in pools of 10 eggs in 1.5 ml tubes. Each tube was spiked with 1,000 P. neurophilia spores and half were treated with RNase AWAY and processed as described above with the remainder untreated. Pseudoloma neurophilia

DNA was detected in the spiked, untreated eggs whereas no P. neurophilia DNA was

detected by qPCR in spiked egg samples that had been treated with RNase AWAY. In

order to determine whether this treatment was affecting DNA present within the egg, qPCR was performed using primers which amplify a fragment of the zebrafish pou5f1 gene (Murray et al., 2011) on eggs which had been treated with RNase AWAY and eggs which had not. Crossing threshold (Cq) values were analyzed using Welch’s t test in the statistical software package R and found to not differ significantly between RNase AWAY-treated and untreated eggs (p > 0.1). DNA extracted from egg pools and spawn water filters was analyzed by qPCR on an ABI 7500 sequence detection system using the method previously described (Sanders and Kent, 2011). For quantification of parasite in samples testing positive, two standard curves were obtained by spiking spawn water filters and egg pools obtained from a group spawn of known Pseudoloma neurophilia-free zebrafish with 100,000 P. neurophilia

spores. Serial two-fold dilutions were made using DNA extracted from these spiked samples and the quantity of parasite was determined by plotting the Cq value of samples against the standard curve. Estimation of P. neurophilia prevalence from pooled samples obtained from paired spawns was calculated using the statistical software environment R and the functions “llprevr” and “dprev” developed by Williams and Moffitt (2005) and is available online (http://www.webpages.uidaho.edu/~chrisw/research/prevalence/). This method calculates pathogen prevalence using a maximum likelihood estimator based on the results of tests performed on samples consisting of variable pool sizes and provides 95% confidence intervals. ACKNOWLEDGEMENTS We thank the Oregon State University Veterinary Diagnostic Laboratory for histological slide preparation and S. Giovannoni for helpful comments and review of this manuscript. This study was supported by grants from the National Institutes of Health (NIH NCRR 5R24RR017386-02 and NIH NCRR P40 RR12546-03S1).

Figure 4.1. Spores of Pseudoloma neurophilia in Luna-stained histological sections of progeny of infected zebrafish, Danio rerio. A. Spores (red) in the epidermis of a 7 d post-fertilization (pf) larval zebrafish. B. Spores in the resorbing yolk-sac of the same 7 dpf larval zebrafish. C. Spore aggregate beneath the intestinal epithelium of an 8 wk pf juvenile fish. D. Spores in the ovigerous stroma adjacent to developing follicles in an 8

wk pf fish.. Bar = 10 µm.

Figure 4.2. Spores of Pseudoloma neurophilia in developing embryo of zebrafish, Danio rerio. A. Aggregated spores (arrow) in a 4 hpf embryo. Bar = 0.5 mm. B. Two foci of spores (arrows) visible in the same embryo at 24 hpf.. C. Spores (arrows) in the same embryo at 48 hpf. D. Differential interference contrast micrograph of spores from an embryo. Bar = 10 µm.

Figure 4.3. Modes of transmission of Pseudoloma neurophilia in the zebrafish, Danio rerio. A. Luna-stained histological section showing P. neurophilia spores (red) within a secondary oocyte. Sexually mature female fish have been shown to harbor the parasite in both ovigerous stromal tissue and within various developmental stages of oocytes. B. Luna-stained histological section of kidney from an adult male zebrafish with spores present (red) within the epithelium of a renal tubule. The presence of spores in these structures is proposed to be one method by which spores can be released into the environment by live fish. C. P. neurophilia spores present within a developing embryo. D. The presence of high numbers of P. neurophilia spores in spawn water and the high susceptibility of larval fish can result in infected progeny. E. Intraovum transmission of P. neurophilia is proposed to result in either the death of the developing embryo or larvae with the subsequent release of spores into the water infecting tank mates or F. live, infected animals that then go on to transmit the parasite horizontally and, later, vertically.

Table 4.1. Detection of Pseudoloma neurophilia in egg pools obtained from paired spawning of Danio rerio adults and tested by qPCR and in spawning adult fish by histology. Estimation of prevalence of P. neurophilia in individual populations of eggs obtained by paired spawning of zebrafish. Prevalence is calculated using the maximum likelihood estimation method of Williams and Moffit (2005). ND = None detected, S= spinal, F = follicle, I = intestinal epithelium, K = kidney, O = ovigerous stroma, P = pancreas. Number Estimated percent positive Number of spores prevalence (95% Spores Total Pool pools/Total detected in confidence detected Histology Histology Spawn eggs Size pools positive egg pools intervals) (water) - Male - Female 1 64 10 0/7 - - 12159 ND S 2 445 30 0/15 - - ND ND ND 3 543 30 0/19 - - 16788 ND S, F 4 463 30 1/16 132560 0.2 (0.01-1.0) ND ND S 5 151 30 2/5 430847, 568 1.3 (0.2-4.0) 1418 ND S 6 107 20 0/6 - - 4417 S, K S 7 223 20 0/12 - - 1858 S S 8 185 30 0/10 - - 12642 S S, O 9 135 20 0/7 - - ND ND S 10 159 20 0/8 - - 1180 S ND 11 295 30 0/11 - - 1385 S S, F 12 323 30 0/12 - - 977 S S 13 72 30 0/3 - - 1549 S, K S, K 14 249 30 0/9 - - ND S ND

15 247 30 0/9 - - 703 S S, I

16 370 20 0/18 - - ND S, K S

17 232 15 0/15 - - ND S ND 18 223 30 0/8 - - ND S S 19 285 30 0/10 - - ND ND S 80

Table 4.1. Continued Number Estimated percent positive Number of spores prevalence (95% Spores Total Pool pools/Total detected in confidence detected Histology Histology Spawn eggs Size pools positive egg pools intervals) (water) - Male - Female 20 193 30 0/7 - - ND S, K S, P 21 177 30 0/6 - - ND S, K S, O 22 76 15 1/6 998 1.5 (0.01-6.3) ND S, K S, O 23 90 30 0/4 - - ND S, K S 24 188 30 1/7 22901 0.6 (0.03-2.5) ND S, K S, K, O 25 178 20 0/9 - - ND S, P S, K 26 154 15 0/10 - - ND S, K S 27 398 30 0/14 - - ND S S,O

Table 4.2. Dose response of larval zebrafish exposed to low numbers of Pseudoloma neurophilia. Two trials were performed in which three replicates of larval zebrafish were exposed to low concentrations of P. neurophilia spores. Fish surviving to seven day post exposure were euthanized and examined by microscopy for the presence of P. neurophilia spores.

500 300 100 50 sp/ml sp/ml sp/ml sp/ml Trial 1

A 5/11 1/14 0/8 0/6 B 2/10 0/9 0/5 0/3 C 6/20 1/11 0/5 0/6 Total percent infected 31.7 5.9 0.0 0.0 Trial 2

A 0/3 1/2 0/4 0/1 B 4/6 1/4 0/4 0/5 C 2/5 1/2 0/1 0/3 Total percent

infected 42.9 37.5 0.0 0.0

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Chapter 5

Early development and tissue distribution of Pseudoloma neurophilia in the zebrafish, Danio rerio

Justin L. Sanders1, Tracy S. Peterson2, Michael L. Kent1,3

Manuscript in preparation

1Department of Microbiology, Oregon State University, Corvallis, OR 97331, USA 2Aquaculture/Fisheries Center, University of Arkansas Pine Bluff, Pine Bluff, AR, 71601, USA 3Department of Biomedical Sciences, Oregon State University, Corvallis, OR 97331, USA

ABSTRACT The early proliferative stages of the microsporidian parasite, Pseudoloma neurophilia were visualized in larval zebrafish, Danio rerio, using histological sections with a combination of an in situ hybridization probe specific to the P. neurophilia small- subunit ribosomal RNA gene, standard hematoxylin-eosin stain, and the Luna stain to visualize spores. Beginning at 5 days post fertilization, fish were exposed to 1.5 x 106 parasites per 100 ml of water and examined at 12, 24, 36, 48, 72, 96, and 120 hours post exposure (hpe). At 12 hpe, P. neurophilia was mainly visualized as intact spores in the intestinal lumen and proliferative stages developing in the epithelial cells of the anterior intestine and the pharynx and within the hepatocytes of the liver. Proliferative stages were visualized in these tissues and in the pancreas and kidney at 36 and 48 hpe and in the spinal cord, eye, and skeletal muscle beginning at 72 hpe. The first spore stages of P. neurophilia were observed at 96 hpe in the pharyngeal epithelium, liver, spinal cord and skeletal muscle. The parasite was only observed in the brain of larval fish at 120 hpe. The distribution of the early stages of P. neurophilia and the lack of mature spores until 96 hpe indicates that the parasite gains access to organs distant from the initial site of entry, likely by penetrating the intestinal wall with the polar tubule, and

that autoinfection does not occur at any detectable frequency during the initial stages of

infection.

INTRODUCTION The microsporidium, Pseudoloma neurophilia, is an obligate intracellular parasite that infects the zebrafish, Danio rerio. The parasite generally results in a chronic infection of adult fish, with spore stages generally found in the anterior spinal cord and nerve root ganglia (Kent and Bishop-Stewart, 2003; Matthews et al., 2001). Subclinical infections of zebrafish are problematic due to the potential for non-protocol induced variation when using infected fish in research (Kent et al., 2012). While much is known about the parasite distribution during later stages of infection, very little is known about the initial stages and, more importantly, how the parasite is able to reach immune- privileged sites such as the spinal cord. In subclinically infected adult fish, P. neurophilia is most commonly observed in immune-privileged sites such as the spinal cord, ventral nerve roots, and anterior brain (Matthews et al., 2001), however, free spores are also often seen in the kidneys and ovaries with the use of chitin-binding fluorescent stains such as Fungi-Fluor (Kent and Bishop-Stewart, 2003). The use of special stains such as Fungi-Fluor and the Luna stain (Peterson et al., 2011) have also enabled the visualization of spores in other tissues, most notably the skeletal muscle of fish with clinical infections due to severe myositis (Kent

and Bishop-Stewart, 2003) and in the ovigerous stroma and within the developing ova of

healthy-appearing females (Sanders et al., 2012). While these special stains provide more sensitive detection of the spore stages of microsporidia in tissues, the visualization of prespore stages of these parasites is much more difficult. In situ hybridization techniques have been used to detect presporogonic stages of microsporidian parasites in a few fish species such as Glugea plecoglossi in rainbow trout (Lee et al., 2004), an unknown species in amberjack (Miwa et al 2011), and Loma salmonae in rainbow trout (Sanchez and Speare, 2001). Sanchez and colleagues used this technique to track the initial stages of the parasite, finding proliferative stages of the parasite in the cells underlying the endocardium, which was present prior to the appearance of xenomas containing mature spores in the gills of infected fish (Sanchez and Speare, 2001).

We infected newly hatched larval fish with P. neurophilia and tracked the infection at various time points post exposure. With the small size of the larvae, we were able to visualize all organs throughout the infection in whole-body coronal sections stained with either hematoxylin and eosin (HE), the Luna stain, or our in situ probe based on the small subunit rDNA gene of the parasite. METHODS Exposures were performed using AB line fish obtained from the P. neurophilia specific pathogen free (SPF) colony located in the Sinnhuber Aquatic Research Laboratory (SARL) at Oregon State University (Kent et al., 2011). Embryos were held in sterile system water at 28° C and checked twice daily. At 5dpf, fish were divided into two separate 250 ml glass beakers in 100 ml of sterile system water each and fed concentrated paramecia twice daily. Spores of P. neurophilia were collected using the method previously described (Ramsay et al., 2009). Briefly, adult fish infected with P. neurophilia were killed by an overdose of tricaine methanesulfate (MS-222), their hindbrains and spinal cords were removed and placed in sterile water containing 100 units each of penicillin and streptomycin (Invitrogen, Carlsbad, CA, USA), and then

macerated by forcing the material through sequentially smaller gauge needles attached to

a 5 ml syringe. Spores were then quantified using a hemocytometer and added to one

6 beaker of larval fish at a concentration of 1.5 x 10 / 100 ml. Larvae in the remaining beaker were maintained as an unexposed control group. In a study of the initial developmental stages of Loma salmonae in rainbow trout, Sanchez et al (2001) first observed intracellular parasite DNA beginning at 12 hours post exposure (hpe). A preliminary study in which we examined larval zebrafish at 1 and 6 hpe confirmed the presence of only extracellular spores in the gut lumen. Thus, exposed larval fish were collected at the following time points in hours post exposure: 0, 12, 24, 36, 48, 72, 96, and 120. Collected larval fish were euthanized by inducing instantaneous fatal hypothermia (ice bath immersion), immediately placed in Dietrich’s fixative and fixed overnight at 4° C. After fixation, embryos were placed in 70% ethanol and embedded in 7 x 4 agarose arrays (Sabaliauskas et al., 2006). The arrays were processed

for histology, paraffin-embedded, and 5 µm serial sections cut with alternating sections stained with HE or Luna, and unstained sections which were examined using in situ hybridization. In situ hybridization. Two oligonucleotide probes previously developed for a real-time PCR based assay (Sanders and Kent, 2011) and specific to the Pseudoloma neurophilia small subunit ribosomal RNA gene were used: P10F-GTAATCGCGGGCTCACTAAG and P10R GCTCGCTCAGCCAAATAAAC. These oligonucleotides were labeled with digoxigenin (DIG) using the 3’ DIG Oligonucleotide Tailing Kit (Roche Applied Science, Indianapolis, IN, USA) following the kit protocol. Tissue sections were deparaffinized by three 10 minute washes in xylene followed by a 3 minute wash in 100% ethanol and rehydration by sequential 3 min washes in progressively lower concentration ethanol solutions (95%, 80%, 70%, 50%) and then 3 min in deionized

water. Tissue sections were then washed in Tris-CaCl3 buffer for 3 min and permeabilized by incubating with proteinase K in Tris-CaCl3 buffer (50 µg/ml) for 15 min at 37°C. After permeabilization, the sections were washed three times in phosphate

buffered saline (PBS) for 10 min each.

Prehybridization was performed by incubating the tissues at 37°C in hybridization

solution (100 µl 20x saline-sodium citrate buffer, 10 µl salmon sperm, 5 mg dextran sulfate, 50x Denhardt’s, 250 µl deionized formamide) without the addition of the digoxigenin-labeled probes for two hours. After two hours, the prehybridization solution was poured off and 60 µl of hybridization solution with 500 ng digoxigenin labeled probes was added to each tissue section. The slides were covered with Hybri-Slips (Sigma-Aldrich, St. Louis, MO, USA), denatured for 10 min at 95°C, and then incubated overnight at 37° C in a MicroProbe slide heater (Fisher Biotech, Fair Lawn, NJ, USA). After incubation, stringency washes were performed using 2 30 minute washes in 2x saline-sodium citrate (SSC, Sigma-Aldrich, St. Louis, MO, USA) buffer at 37°C, 3 10 minute washes in 1x SSC at 37°C , and one 10 minute wash in 0.5x SSC at room temperature. Following the stringency washes, the tissue sections were washed in Wash

Buffer (Roche Applied Science, Indianapolis, IN, USA) for 10 min at room temperature and then soaked in maleic acid Blocking Buffer (Roche Applied Science, Indianapolis, IN, USA) for one hour at room temperature. The sections were then incubated for 2 hours with anti-DIG antibody (Roche Applied Science, Indianapolis, IN, USA) diluted 1:1000 in Blocking Buffer at room temperature. The antibody solution was then poured off and the slides were washed twice for 15 min each in Wash Buffer on a shaker. They were then soaked in Detection Buffer (Roche Applied Science, Indianapolis, IN, USA) for 10 min after which they were drained and substrate, nitroblue tetrazolium NBT/BCIP Ready-to-Use Tablets (Roche Applied Science, Indianapolis, IN, USA) dissolved in Detection buffer was added for 1-2 hours. After examination that the blue reaction had occurred, the slides were washed twice in deionized water. Tissue sections were counter-stained using Nuclear Fast Red (Vector Laboratories, Inc, Burlingame, CA, USA) for 5 min, followed by rinsing in deionized water and air drying. The tissue sections were then dehydrated by subsequent washing in 70% ethanol for 3 min, 95% ethanol for 3 min, and two changes of 100% ethanol for 3 min each. Finally, the tissue sections were soaked in two changes of xylene for 3 min

each and coverslipped using Cytoseal XYL (Richard Allan Scientific, Kalamazoo, MI,

USA) permanent mounting medium.

RESULTS Chronological sequence of Pseudoloma neurophilia progressive infection in the larval zebrafish is presented in Table 5.1. Occasional intraluminal loose aggregates of individual mature intact spores were observed within the anterior intestine by Luna stain at 12 hours post-exposure (hpe), likely reflecting ingestion by larval fish (Fig. 5.1). The observation of mature spores within the intestinal lumen declined during the later time points and was no longer observed after 72 hpe. The initial observation of mature spores developing within host tissues was noted at 96 hpe. No parasites were observed in the unexposed negative control fish.

Intestine Presporogonic proliferative stages of P. neurophilia were initially observed in the digestive tract at 12 hpe in both the pharyngeal (Fig. 5.2) and intestinal epithelia (Figs. 5.3, 5.4) by in situ hybridization and Luna stain. Developing spores were localized within the apical cytoplasmic compartment of infected cells and tended to be found in the anterior segment of the intestine. Proliferative stages of the parasite continued to be observed in fish collected from all later time points; however, mature spore stages were only observed in the pharyngeal epithelium at 96 hours post-exposure (hpe) and in the intestinal epithelium at 120 hpe. Visceral organs and kidney Within the liver (Fig 5.5a, 5.5b), intrahepatocytic presporogonic developmental stages of P. neurophilia were initially observed at 12 hpe by in situ hybridization (Fig. 5.6) followed by the appearance of mature spores at 96 hpe, which were detected by the Luna stain. Beginning at 36 hpe, similar presporogonic proliferative stages of P. neurophilia were observed associated with endothelial cells lining an intrapancreatic blood vessel (Fig. 5.7) and intracellular mature spores were seen in pancreatic acinar cells

of a larval fish at 120 hpe. Presporogonic developmental stages involving the kidney

were confined to occasional histiocytes within the renal interstitium (Fig. 5.8) beginning

at 48 hpe. Muscle and neural Presporogonic stages of P. neurophilia were first observed in the spinal cord, skeletal muscle, and eye at 72 hpe. In the spinal cord, small aggregates of presporogonic stages were distributed among ependymal cells forming the lining of the central canal as highlighted by in situ hybridization (Fig. 5.9). Mature spores were observed in the spinal cord 96 hpe. Intrasarcolemmal dense aggregates of proliferative stages were observed within individual myofibers of skeletal muscle (Fig. 5.10), with mature spores first observed 96 hpe. Within the extraocular choroid rete, small aggregates of P. neurophilia merogonic stages were observed in the non-vascular stroma immediately adjacent to the retinal pigmented epithelium (Figs. 5.11-5.13) and within the retinal pigmented

epithelium, extending into the photoreceptive layer (Figs. 5.14-5.16), by in situ hybridization and Luna stain. Mature spores were observed in these locations at 120 hpe. Brain neuropil contained both proliferative and mature spore stages (Figs. 5.17, 5.18), which were observed at 120 hpe only. DISCUSSION Determining the mechanisms of initiation of infection and the distribution of parasites within the host in the early stages of microsporidian infections is important to the understanding of systemic microsporidiosis. Using HE and Luna stains in combination with ISH enabled the observation of the earlier, presporogonic stages of Pseudoloma neurophilia and its distribution in tissues. Additionally, the use of larval zebrafish enabled us to examine several individual whole animals on a single slide. By performing serial sections, virtually all organs of each animal were examined. This method allows for the comprehensive analysis of the early development and tissue progression of P. neurophilia in the larval zebrafish, therefore expanding the understanding of initial infection and parasite distribution and development beyond our previous studies (Cali et al., 2012; Kent and Bishop-Stewart, 2003; Sanders et al., 2012).

The following summarizes our understanding of the sequential development of P.

neurophilia: Spores are ingested and germinate in the anterior intestine. By 12 hpe

presporogonic proliferative stages are observed in the intestinal and pharyngeal epithelia, and liver. Beginning at 36 hpe, presporogonic proliferative stages are found in the pancreas, and shortly thereafter in the kidney. At 72 hpe, presporogonic proliferative stages are first seen in the spinal cord, eye, and skeletal muscle. The first time developed spores are observed is at 96 hpe in the visceral organs, followed shortly thereafter in the CNS and the skeletal muscle. It is well-recognized that microsporidia initiate infection of host cells by extrusion of their polar tube and infection of the sporoplasm into host cells (Cali and Takvorian, 1999). Following ingestion, spores adhere to gastrointestinal epithelia associated with sulfated glycans (Hayman et al., 2005). Spores may then extrude their polar tube and infect adjacent intestinal cells. Alternatively, spores may be phagocytosed

by host cells in the gut, then extrude their polar and infect the same host cell (Couzinet et al., 2000). Polar tubes are range in length from 50-100 µm, and Cox et al. (Cox et al., 1979) proposed a third mechanism; injection of the polar tube through the intestine to more distant tissues. Pseudoloma neurophilia initially infects the host by ingestion of the infective spore stage, with spores being observed in the gut lumen of exposed larval fish at 3 hpe (Cali et al., 2012). Merogonic and sporogonic stages can be observed in the skeletal muscle at 108 hpe (Cali et al., 2012). That observation was confirmed by the current study in which the first stages observed in skeletal muscle were found at 72 hpe. In addition, we found numerous other tissues that were infected shortly after exposure, notably, the pancreas, liver, kidney and the extraocular tissues and retina of the eye. However, we did not observe merogony in the intestinal epithelium or underlying lamina propria. Hence, our study supports the mechanism purposed by Cox et al. (1979), piercing of the intestinal wall by the polar tube to infect distant tissues. The sites of initial parasite development that we observed are within the range of the polar tubule, which is greater than 100 µm in length. This indicates that the spore germinates with the apical cap oriented facing the intestinal epithelium, firing the polar tubule and acting as a

syringe to penetrate the intestinal wall and infect distant tissues, such as the liver or

pancreas, and injects the sporoplasm at these sites. Indeed, far more developing parasites were observed in the liver, kidney and pancreas during early stages of infection, rather than within the intestinal tissues. Hayman and colleagues (2005) showed that Encephalitozoon intestinalis spores bind to sulfated glycans on the surface of host cells and that this adherence was important to the infectivity of those spores. There is some evidence to support this such as the specificity of germination triggers possessed by different species of the Microsporidia. The tissue tropism of a particular microsporidian species could be controlled by the environmental cue for germination (usually in the gastrointestinal tract), resulting in a spore firing only when this cue is sensed. The exact trigger for P. neurophilia is unknown and we have never observed firing of the polar tubule except when spores are treated with a highly alkaline, chitin binding stain (Fungi

Fluor) and exposed to the UV light of a fluorescence microscope (Ferguson, Watral, Schwindt, & Kent, 2007), a situation not encountered within live zebrafish tissues. The tight control of spore germination by a mechanism such as adhesion to host surface factors would prevent or limit unsuccessful infections by spores. In an in vitro study of the early development of the microsporidium, Anncalia algerae, in rabbit kidney cells, Takvorian et al. (2005) did not observe mature spore formation in 48 hpe cells, but they did observe intracellular sporoplasms and early stages of the parasite in cell cultures incubated for up to 48 hours, suggesting that these were new infections. They attributed these new infections observed several hours post inoculation to delayed spore germination and suggested that delayed spore activation was possibly an adaptation, allowing a population of parasites to infect various sites of the host (Takvorian et al., 2005). As this observation was made in cultured cells, this is likely true in their study. Although we observed presporogonic stages several days after the initial exposure, these were in tissues distant from the intestinal epithelium. There could be a number of mechanisms responsible for this observation, such as the transport of the parasite within a motile host cell (e.g., a macrophage), or the piercing of the intestinal

epithelium by the polar tubule of the parasite and the injection of the sporoplasm directly

into the blood or the cytoplasm of the host cell in which the earliest stages of the parasite

were observed As the name implies, P. neurophilia, is most often found in the neural tissue, mainly the ventral nerve root ganglia, metencephalon and myelencephalon (together comprising the hindbrain) of chronically infected zebrafish. Kent and Bishop-Stewart (2003) performed a histological survey of the tissue distribution of P. neurophilia in adult zebrafish and compared the distribution between subclinical and clinically infected fish. Using a chitin-specific fluorescent stain, Fungi-Fluor, they were able to increase the sensitivity of detection of the spore stage of the parasite in tissue sections over the use of the standard HE stain. Peterson and colleagues (2011), found that the use of the Luna stain similarly increased the sensitivity of the detection of spores in histological sections without the need for fluorescence microscopy.

Whereas the parasite is seen in the skeletal muscle in the early stages of infection, in chronic infections of ostensibly immunocompetent zebrafish hosts, P. neurophilia is generally isolated in immune-privileged sites such as the spinal cord, hindbrain and developing ova (Matthews et al., 2001). We observed P. neurophilia proliferative stages in the spinal cord and eye as early as 72 hpe and in the brain at 120 hpe. The observation of P. neurophilia developing in the choroid rete and pigmented retinal epithelia of the eyes of several zebrafish is a heretofore unreported site of infection for P. neurophilia. Other microsporidian species infecting humans have been documented to cause ocular infections (Friedberg and Ritterband, 1999). In immunocompetent patients, these infections generally occur deep in the corneal stroma, occasionally associated with prior trauma, and are not associated with systemic microsporidiosis (Weber et al., 1994). In immunosuppressed patients, while infections are generally limited to the superficial epithelium of the cornea, they are often associated with systemic infection (Weber et al., 1994). The exact mechanism of the movement of P. neurophilia within the body of the zebrafish after initiation still not completely elucidated, as was the case with experimental infection studies with other systemic microsporidia (Cox et al., 1979).

Sanchez et al. (2001) used in situ hybridization to describe the development of the

microsporidium, Loma salmonae in the Atlantic salmon, Salmo salar, and found that shortly after infection, which begins in the intestinal epithelium, merogonic stages can be seen within the intertrabecular spaces of the ventricular spongy myocardium of the heart and along the endocardial lining of the ventricular trabeculae at 2 dpe. Consistent with our findings, Sanchez et al. (2001) first observed proliferative stages of Loma salmonae in the intestinal epithelium at 12 hpe. Whereas we observed mature (Luna-positive) spores of P. neurophilia in various tissues as early as 96 hpe, the first spores of L. salmonae were observed at 4 wk post exposure and localized to the gills (Rodríguez- Tovar et al., 2003). The authors hypothesized that the parasite moved from the intestinal epithelium to the heart by infecting mobile leukocytes, such as monocytes. As the endocardial cells in the heart function as phagocytic cells (i.e., macrophages) in teleost

fishes, and these cells are maybe “grabbing” and sequestering the parasite as it enters systemic circulation. Both Loma and Pseudoloma have been observed within macrophages, adding support to this hypothesis. The use of real-time live imaging of an infection of a larval zebrafish by labeled P. neurophilia would likely enable us to definitively determine the mode of transport. Unfortunately, current lack of tools to produce transgenic microsporidia prevents this type of experiment. In conclusion, we expanded our understanding of the early development, organ distribution, timing and location of sporulation of P. neurophilia in larval zebrafish. Most notably we observed first sporulation concurrently in the visceral organs and the CNS, whereas the latter has been previously considered the primary site of infection. Additionally, we have observed for the first time the parasite developing in the choroid rete and pigmented retinal epithelium of the eye. The retina is an extension of the central nervous system, thus is consistent with the neurotropism of this microsporidium. Both larval and post-larval fish are susceptible to natural transmission of the parasite (Ferguson et al., 2007; Kent and Bishop-Stewart, 2003; Sanders et al., in press), including maternal transmission to embryos and fry (Sanders et al., in press). In chronic infections of older fish, P. neurophilia is most commonly found in immune-privileged sites such as the

spinal cord, nerve root ganglia, hindbrain, and occasionally developing ova. The eye

could be another target site for latent infection. A comparison of our findings in larval zebrafish to the early stages of progression and development of P. neurophilia in juvenile or adult fish is warranted. ACKNOWLEDGEMENTS We thank the Oregon State University Veterinary Diagnostic Laboratory for histological slide preparation. This study was supported by grants from the National Institutes of Health (NIH NCRR 5R24RR017386-02 and NIH NCRR P40 RR12546-03S1).

100

Fig 5.1-5.4. Early stages of Pseudoloma neurophilia infection in the intestinal tissue of larval zebrafish at 12-72 hours post-exposure. Bars = 10 µm. 1. Luna-stained section of a larval fish after 12 hours post-exposure to P. neurophilia. Several individual mature intact spores (red) are visible in the lumen of the anterior intestine. 2. Section of a larval fish after 36 hours post-exposure to P. neurophilia stained using an in situ hybridization

probe (ISH) specific to P. neurophilia. Presporogonic proliferative stages are visible (arrow) developing in the pharyngeal epithelium. 3. ISH stained section of a larval fish

after 48 hours post-exposure. A single proliferative stage is visible (arrow) in the cytoplasm of an intestinal epithelial cell. 4. Hematoxylin and eosin stained section of a larval zebrafish at 72 hours post-exposure. Presporogonic proliferative stages in the cytoplasm of an intestinal epithelial cell (arrow).

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Fig. 5.5-5.8. Early stages of Pseudoloma neurophilia infection in extraintestinal organs of larval zebrafish. Bars = 10 µm. 5a. Hematoxylin and eosin stained section showing the liver of a larval zebrafish at 72 hours post-exposure. Nucleated erythrocytes (e), hepatocyte nuclei (h) and a capillary (c) can be observed. 5b. High magnification of the boxed area in 5a. A cluster of presporogonic proliferative stages (arrow) can be seen developing in a hepatocyte. Note proximity to capillary (C). 6. Section of a larval zebrafish at 72 hours post-exposure, stained with an in situ hybridization (ISH) probe specific to P. neurophilia. Presporogonic proliferative stages (arrow) developing within a hepatocyte. 7. ISH stained section of a larval zebrafish at 72 hours post-exposure. Three presporogonic proliferative stages (arrows) associated with endothelial cells lining an intrapancreatic blood vessel . 8. ISH stained section of a larval zebrafish at 72 hours post-exposure. A single presporogonic proliferative stage (arrow) developing within the cytoplasm of a kidney histiocyte.

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Fig. 5.9-5.18. Early stages of Pseudoloma neurophilia in neural tissues of larval zebrafish. Bars = 10 µm. 9. Section of a larval zebrafish at 5 days post-exposure stained

with an in situ hybridization (ISH) probe specific to P. neurophilia. Presporogonic proliferative stages (blue) and spores (arrow) developing among ependymal cells lining

the central canal of the spinal cord. 10. ISH stained section of a larval zebrafish at 5 days post-exposure. A dense aggregate of proliferative stages (blue) developing within an individual myofiber. 11. ISH stained section of a larval zebrafish at 72 hours post- exposure. Proliferative stages and spores (arrow) within the extraocular choroid rete adjacent to the retinal pigmented epithelium. 12. Adjacent section of the previous fish stained with the Luna stain. Red staining mature spores (arrow) are more apparent within the extraocular choroid rete. 13. Adjacent section of the previous fish stained with hematoxylin and eosin (HE). Mature spores (arrow) are faintly visible in the extraocular choroid rete. 14. Adjacent section of the previous fish stained with ISH. Proliferative stages are visible (arrow) in the retinal pigmented epithelium extending into the photoreceptive layer. 15. Adjacent section of the previous fish stained with the Luna stain. Red-staining mature spores (arrow) are visible in the retinal pigmented epithelium. 16. Adjacent section of the previous fish stained with HE. No presporogonic nor spore stages are visible. 17. ISH stained section of a larval zebrafish at 5 days post-exposure. A proliferative stage is visible (arrow) developing within the brain neuropil. 18. An HE stained section of a larval zebrafish 5 days post-exposure. A cluster of proliferative stages is visible within the brain neuropil.

Table 5.1. Results of histological examination of larval zebrafish at various times post exposure to Pseudoloma neurophilia. Larval zebrafish were exposed to P. neurophilia spores and collected at 12-120 hours post exposure. HPE = hours post exposure, P = presporogonic proliferative stages, S = spore stages.

Total fish Intestinal Intestinal Spinal HPE examined lumen epithelium Pharynx Liver Pancreas Kidney Cord Eye Muscle Brain P S P S P S P S P S P S P S P S P S P S 12 28 0 20 6 0 3 0 4 0 0 0 0 0 0 0 0 0 0 0 0 0 24 28 0 9 4 0 2 0 3 0 0 0 0 0 0 0 0 0 0 0 0 0 36 28 0 6 8 0 5 0 4 0 4 0 0 0 0 0 0 0 0 0 0 0 48 28 0 3 12 0 9 0 12 0 0 0 2 0 0 0 0 0 0 0 0 0 72 28 0 1 11 0 13 0 17 0 6 0 4 0 1 0 3 0 8 0 0 0 96 21 0 0 2 0 3 2 3 2 2 0 0 0 1 1 0 0 1 1 0 0 120 22 0 0 8 2 4 1 15 7 7 1 13 0 7 3 12 9 17 14 4 4

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REFERENCES Cali, A., and Takvorian, P. (1999). Developmental morphology and life cycles of the Microsporidia. M. Wittner, and L.M. Weiss, eds. (Washington DC: ASM Press), pp. 85– 128.

Cali, A., Kent, M., Sanders, J., Pau, C., and Takvorian, P.M. (2012). Development, ultrastructural pathology, and taxonomic revision of the microsporidial genus, Pseudoloma and its type species Pseudoloma neurophilia, in skeletal muscle and nervous tissue of experimentally infected zebrafish Danio rerio. J. Eukaryot. Microbiol. 59, 40– 48.

Couzinet, S., Cejas, E., Schittny, J., Deplazes, P., Weber, R., and Zimmerli, S. (2000). Phagocytic uptake of Encephalitozoon cuniculi by nonprofessional phagocytes. Infect. Immun. 68, 6939–6945.

Cox, J.C., Hamilton, R.C., and Attwood, H.D. (1979). An investigation of the route and progression of Encephalitozoon cuniculi infection in adult rabbits. J. Eukaryot. Microbiol. 26, 260–265.

Friedberg, D.N., and Ritterband, D.C. (1999). Ocular microsporidiosis. In The

Microsporidia and Microsporiosis, M. Wittner, and L.M. Weiss, eds. (Washington, D.C.: ASM Press), pp. 293–313.

Hayman, J.R., Southern, T.R., and Nash, T.E. (2005). Role of sulfated glycans in adherence of the microsporidian Encephalitozoon intestinalis to host cells in vitro. Infect. Immun. 73, 841–848.

Kent, M.L., and Bishop-Stewart, J.K. (2003). Transmission and tissue distribution of Pseudoloma neurophilia (Microsporidia) of zebrafish, Danio rerio (Hamilton). J. Fish Dis. 26, 423–426.

Kent, M.L., Harper, C., and Wolf, J.C. (2012). Documented and potential research impacts of subclinical diseases in zebrafish. ILAR J. Natl. Res. Counc. Inst. Lab. Anim. Resour. 53, 126–134.

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Lee, S.-J., Yokoyama, H., and Ogawa, K. (2004). Modes of transmission of Glugea plecoglossi (Microspora) via the skin and digestive tract in an experimental infection model using rainbow trout, Oncorhynchus mykiss (Walbaum). J. Fish Dis. 27, 435–444.

Rodríguez-Tovar, L.E., Wright, G.M., Wadowska, D.W., Speare, D.J., and Markham, R.J.F. (2003). Ultrastructural study of the late stages of Loma salmonae development in the gills of experimentally infected rainbow trout. J. Parasitol. 89, 464–474.

Sanchez, J.G., and Speare, D.J. (2001). Localization of the initial developmental stages of

Loma salmonae in rainbow trout (Oncorhynchus mykiss). Vet. Pathol. 38, 540–546.

Sanders, J.L., Watral, V., Clarkson, K., and Kent, M.L. (in press). Verification of intraovum transmission of vertebrates: Pseudoloma neurophilia infecting the zebrafish, Danio rerio. PLoS ONE.

Sanders, J.L., Watral, V., and Kent, M.L. (2012). Microsporidiosis in zebrafish research facilities. ILAR J. Natl. Res. Counc. Inst. Lab. Anim. Resour. 53, 106–113.

Takvorian, P.M., Weiss, L.M., and Cali, A. (2005). The early events of Brachiola algerae (Microsporidia) infection: spore germination, sporoplasm structure, and development within host cells. Folia Parasitol. (Praha) 52, 118–129.

Weber, R., Bryan, R.T., Schwartz, D.A., and Owen, R.L. (1994). Human microsporidial infections. Clin. Microbiol. Rev. 7, 426–461.

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Chapter 6

The Zebrafish as a Model for Microsporidiosis

Justin L. Sanders1, Michael L. Kent1,2

Microsporidia: Pathogens of Opportunity, In Press Wiley-Blackwell Publishing, Hoboken, NJ, USA

1Department of Microbiology, 220 Nash Hall, Oregon State University, Corvallis, Oregon, 97331 2Department of Biomedical Sciences, Oregon State University, Corvallis, Oregon, 97331

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INTRODUCTION The zebrafish, Danio rerio, is now one of the most important vertebrate models in biomedical research, and can be naturally infected with two microsporidia. Therefore, this fish provides an excellent platform for investigating various basic and applied research topics pertaining to microsporidia in vertebrates. Here we review and discuss the zebrafish model and its documented and potential uses in the study of microsporidia. The zebrafish is an important laboratory model for toxicology, developmental biology, cancer, and infectious disease research. It has a well-characterized immune system and the complete genome has been sequenced with the genetic map showing an overall highly conserved synteny with the human genome (Catchen et al., 2011; Howe et al., 2013; Postlethwait et al., 1998). The model was originally widely used for developmental genetics. More recently, juvenile and adult zebrafish have been used in in vivo experiments using various bacterial and viral pathogens to investigate host-pathogen interactions. The results of several infectious disease studies using zebrafish have shown a number of immune response pathways which are analogous to those of humans and other mammals. The high prevalence of a microsporidian parasite in laboratory zebrafish

populations, Pseudoloma neurophilia, provides an ideal opportunity to learn more about

host responses to microsporidian infections using a well-characterized vertebrate model

organism. Additionally, the recent description of another microsporidian infecting zebrafish, Pleistophora hyphessobryconis, provides an opportunity to study the pathological changes associated with two different microsporidian parasites with very distinctive tissue tropisms: P. hyphessobryconis primarily infects skeletal myocytes and P. neurophilia is primarily found in myelinated neural tissue. HISTORY OF ZEBRAFISH IN RESEARCH The zebrafish, Danio rerio (Hamilton-Buchanan, 1822, 1823) is a tropical fish belonging to the family Cyprinidae. Normally inhabiting floodplains surrounding the Ganges and Brahmaputra river basins of northeastern India, the zebrafish has been popular with hobby aquarists for over a century. Zebrafish generally occupy slow- moving waters with a wide range of temperatures (6°C - 38°C) (Spence et al., 2008). The

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wide temperature range and variable water quality parameters found in the natural habitat of the zebrafish explains the hardiness and resilience of this small fish, and makes it attractive as an aquarium species and for research. Zebrafish embryonic development has been investigated in the laboratory since the 1930s (Laale, 1977). In the mid-1950s, zebrafish were used as a bioassay to detect a number of cytotoxic and teratogenic substances in water (Jones and Huffman, 1957). The use of zebrafish as a genetic model for embryogenesis in the laboratory expanded rapidly due to the work begun by Dr. George Streisinger, University of Oregon, in the late 1960s undertaken in order to develop a vertebrate model for the study of development (Grunwald and Eisen, 2002). His reasoning for the selection of the zebrafish for this endeavor were mainly practical, namely: 1. Zebrafish are prolific breeders and produce hundreds of progeny per female in the laboratory setting, 2. Zebrafish embryos are fertilized externally, allowing for the separate harvest of gametes and 3. Embryonic development occurs ex vivo within a transparent egg, allowing researchers to view the entire process. All of these qualities of the zebrafish make it an exceptional vertebrate model for the study for development, as well as for other research

areas, such as infectious diseases.

In the past, researchers would often obtain zebrafish from pet stores. Whereas

this suited the purposes of many investigators at the time, the field has benefitted from the development and maintenance of zebrafish colonies specifically for research purposes. In 1997, the Trans-NIH Zebrafish Initiative was begun in order to further develop zebrafish as a model system for the genetic studies of disease and embryogenesis (Henken, 1998) in response to recommendations by leading zebrafish researchers. In 2000, the Zebrafish International Resource Center (ZIRC) was formed to at the University of Oregon to act as a repository for the maintenance and distribution of zebrafish mutant lines as well as the development of the Zebrafish Information Network (ZFIN), a web-accessible database providing comprehensive information regarding zebrafish, including protocols, for the zebrafish research community (Sprague et al., 2001). In addition, the ZIRC provides a diagnostic and screening service for zebrafish

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researchers to submit samples. Books and review articles are also available, providing a wealth of information covering virtually all aspects of zebrafish husbandry, colony management, and general protocols for the use of zebrafish in research (Harper and Lawrence, 2010; Westerfield, 2007). The zebrafish has recently become a very widely-used model for infectious diseases (Meijer and Spaink, 2011). Research using zebrafish to model fish and human pathogens has provided a wealth of knowledge to the understanding of host-pathogen interactions. Currently, zebrafish models exist for a number of bacterial, viral, and fungal pathogens, including several that infect humans. The list includes Salmonella typhimurium (van der Sar et al., 2003, 2006; Stockhammer et al., 2009), Salmonella arizonae (Davis et al., 2002), Vibrio anguillarum (O’Toole et al., 2004), Citrobacter freundii (Lü et al., 2011, 2012), Bacillus subtilis (Herbomel et al., 2001), Listeria monocytogenes (Levraud et al., 2009), Listeria spp, Streptococcus iniae (Neely et al., 2002), Streptococcus pyogenes (Neely et al., 2002), Streptococcus agalactiae (Patterson et al., 2012), Streptococcus suis (Wu et al., 2010), Mycobacterium marinum (Clay et al., 2007; Davis et al., 2002; Hegedus et al., 2009; Meijer et al., 2005; Tobin et al., 2010),

spring viremia of carp virus (Sanders et al., 2003), infectious haematopoietic necrosis

virus (LaPatra et al., 2000; Ludwig et al., 2011), viral haemorrhagic septicemia virus

(Encinas et al., 2010; Novoa et al., 2006) and Candida albicans (Brothers et al., 2011; Chao et al., 2010), and the list is growing. The presence of two microsporidia that infect zebrafish, Pseudoloma neurophilia and Pleistophora hyphessobryconis, provides an opportunity to study several aspects of microsporidian infection using a well-established vertebrate model system. ADVANTAGES OF THE ZEBRAFISH MODEL From its origins as a model for developmental biology (Streisinger et al., 1981), the use of zebrafish in research has diverged to include toxicology (Spitsbergen and Kent, 2003), drug development (Zon and Peterson, 2005), cancer (Amatruda et al., 2002), hematopoiesis (Davidson and Zon, 2004; Paik and Zon, 2010), immunology (Balla et al., 2010; Traver et al., 2003; Trede et al., 2004), and infectious disease (Meeker and Trede,

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2008; Meijer and Spaink, 2011; van der Sar et al., 2004; Sullivan and Kim, 2008). The same characteristics which make the zebrafish so amenable to developmental research make it highly desirable in these other areas of research, namely, ease of husbandry, high fecundity, small size, availability of numerous lines, and rapid ex vivo development of its transparent embryo. Since emerging as a prominent biological model, numerous other advances have resulted in the rapid spread of the use of zebrafish in research. Sequencing of the genome of the zebrafish began in 2001 and comparison of the latest version of the reference assembly with the human genome has revealed that 70% of human genes have at least one clear zebrafish orthologue (Howe et al., 2013). Current next generation sequencing technology has also allowed for extensive gene transcript sequencing (RNA-Seq) and the inclusion of gene transcript sequences has greatly enhanced the annotation of the zebrafish genome (Collins et al., 2012), providing the basis for the development of molecular tools to study aspects of zebrafish biology. Indeed, these tools provide a number of opportunities for the study of infectious disease processes using the zebrafish model system, as can be exemplified by several recent studies which take advantage of RNA-Seq to examine host gene-expression

changes in response to pathogens (Hegedus et al., 2009; Ordas et al., 2011; Stockhammer

et al., 2010).

Ease of husbandry The zebrafish is a very hardy animal, capable of living comfortably within a wide range of environmental conditions in a various types of aquatic housing systems. While traditional aquaria may be used to house zebrafish for small-scale experiments, research- grade modular and custom rack systems are available from several vendors. Many of these rack units use a recirculating water system in which water is flowed into the tanks continuously or intermittently and effluent from fish tanks is circulated back into the tanks after being treated to remove waste products. The nature of infectious disease research, however, generally requires the use of static or flow-through tank systems in which the effluent is discarded in order to avoid cross-contamination of treatment groups. Zebrafish embryos and larvae can be maintained in static water systems and short-term

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experiments using these stages can easily be set up in 96-well or other sized multi-well plates for the duration of the study (Mandrell et al., 2012; Zon and Peterson, 2005). There are a number of widely available resources for information regarding all aspects of husbandry of zebrafish including books (Harper and Lawrence, 2010; Westerfield, 2007) and internet resources such as the Zebrafish Information Network (ZFIN). The Zebrafish International Resource Center also provides an online manual of common diseases seen in zebrafish research facilities. There are even professional organizations such as the Zebrafish Husbandry Association (www.zhaonline.org), an organization devoted to the promotion and development of standards and education related to laboratory zebrafish husbandry, which are available to provide educational resources. Fecundity Zebrafish are highly fecund and able to produce clutches of up to several hundred eggs with regularity upon reaching sexual maturity, which generally occurs within 3-6 months depending upon nutrition and environmental conditions (Spence et al., 2008). There are several methods and specialized equipment for breeding zebrafish but several

hundred embryos can be obtained using minimal equipment. Laboratory-reared zebrafish

have been shown to spawn on average every 1.9 days (Eaton and Farley, 1974).

Small size: Amenable to high-throughput assays In addition to allowing for a high density of animals with which to do research, the small size of the zebrafish (3 – 5 cm as adults) gives researchers the ability to examine tissues in the context of the entire animal in histological sections. The ability to examine parasites in the context of intact tissue within the body of the organism provides obvious advantages to studies that rely on histopathology. The use of larval fish provides the opportunity to examine numerous individual fish on a single slide (Fig. 6.1). Sabaliauskas et al (Sabaliauskas et al., 2006) described a high-throughput method for larval zebrafish histology. Using 7 dpf fish, this method allows for the visualization of several, in this case 50, fish per section. The ability to visualize many specimens on a

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single slide enables histopathologists to detect subtle changes to tissues in response to microsporidian infection and provides studies with increased rigor. The small size of the larval zebrafish also makes them amenable to studies using electron microscopy. For example, an ultrastructural analysis of the development of P. neurophilia using intact larval fish was performed by Cali et al (Cali et al., 2012), leading to a detailed description of the host-parasite interface. Thus, the zebrafish larva/Pseudoloma model provides an excellent format to investigate initial infection and early parasite development in a vertebrate model. Optical clarity Even as a larva, the zebrafish maintains a great deal of clarity, allowing for direct visualization of host-microsporidium interactions in fish at later stages. The use of in situ hybridization on both whole larval fish and tissue sections has been widely used by developmental biologists and in studies using specific DNA probes to detect pre sporogonic stages of microsporidia in fish tissues (Grésoviac et al., 2007; Sanchez et al., 2001; Yokoyama et al., 2008). In fact, specific in situ DNA probes are currently being used in our laboratory to detect and track pre-sporogonic stages and development of P

neurophilia. The development of zebrafish lines containing transgenic fluorescent

reporter genes allows for the real-time visualization of specific host cell populations in

larval fish. Whereas zebrafish embryos are transparent, the production of pigments in larval fish renders them opaque after a few weeks of development. Thus several lines of zebrafish lacking black melanophores and reflective iridophores that are transparent as adults have been developed, allowing for the visualization of internal organs within adult animals. One such line that is popular with researchers is the casper (White et al., 2008). These fish have been used to examine the engraftment characteristics of transplanted hematopoeitic and tumor cells in live adult fish (Taylor and Zon, 2009; White et al., 2008). As tools for microsporidia, such as mutants and fluorescently labeled organisms, are developed, the transparency of the zebrafish can easily be exploited to study the

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progression of microsporidiosis in vivo. For example, several fluorescent protein expressing transgenic bacterial strains have been visualized in real-time using larval zebrafish, providing insights into basic mechanisms of bacterial pathology such as the formation of granulomas by Mycobacterium marinum (Swaim et al., 2006). The development of a fluorescently labeled microsporidium would allow for the ability to track parasites through whole adult fish by intravital imaging. Availability of numerous lines Numerous transgenic lines of zebrafish have been developed with fluorescent labeling of specific gene products in order to visualize and identify specific cells in vivo. Of particular interest to microsporidian research are the labeling of specific cell populations of the innate immune system such as green fluorescent protein (GFP) labeling of myeloperoxidase (MPO) which specifically labels neutrophils (Renshaw et al., 2006). In addition, numerous lines of zebrafish are available with mutations that affect various parts of the immune system, such as the complete inactivation or deletion of specific cell lineages. The use of such fish in microsporidian research could elucidate the effects of specific immune cells on microsporidian dispersal throughout the body and

pathology. For example, mutant lines lacking specific components of the immune

system, such as those developed that are deficient in rag1 (Wienholds et al., 2002), a

product which is required for functional T and B cells, can be used to study the effects of adaptive immunity on microsporidian infection. The use of animals with underlying chronic infections can result in non-protocol induced variation, confounding many studies. This is especially true when performing infection studies. Pseudoloma neurophilia is commonly found in zebrafish facilities (Murray et al., 2011) and pre-existing infections of fish used to study P. neurophilia, or any other microsporidians, would prove detrimental to research. The growing interest in the use of zebrafish for many types of research and the recognition of the negative impacts of chronic infections on research (Kent et al., 2012) has led to the development of numerous Pseudoloma neurophilia specific-pathogen free (SPF) zebrafish lines (Kent et al., 2011). The low prevalence of Pleistophora hyphessobryconis in laboratory strains

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of zebrafish obviates the need for rigorous screening for this pathogen, however, researchers should be cautioned against obtaining fish from suppliers who deal with multiple species of fish due to the broad host range of this microsporidium. See Kent and colleagues (Kent et al., 2009) for a general review of the control of diseases in fish research colonies. Ex vivo development of transparent embryo As the embryo of the zebrafish is transparent and develops ex vivo, a characteristic which has been widely exploited by developmental biologists, this aspect of the zebrafish can also be useful in the study of microsporidian transmission. For example, the visualization of the microsporidium, Pseudoloma neurophilia, within the yolk of a developing embryo (Fig. 6.2), provides direct, visual evidence that intraovum transmission of this parasite occurs (Sanders et al., in press). In addition, microinjection of microsporidia into embryos at various stages provides one way in which host microsporidian interactions between specific cell types could be easily visualized. DEVELOPMENT/CHARACTERIZATION OF ZEBRAFISH AS A BIOMEDICAL MODEL

Immunology

Extensive work has been done to characterize the ontogeny and function of the

components comprising the zebrafish immune system (Meeker and Trede, 2008; Traver et al., 2003). Zebrafish, as teleost fish, have a fully developed innate and adaptive immune system and share many of the same immune cells and markers with humans (Meeker and Trede, 2008; Traver et al., 2003). Most of the cells comprising cellular immunity have been characterized in the zebrafish and have been found to be highly functionally conserved with mammalian cell types including: macrophages (Herbomel et al., 2001), neutrophils (Balla et al., 2010; Mathias et al., 2009), T cells (Danilova et al., 2004), B cells (Danilova and Steiner, 2002), and dendritic antigen presenting cells (Lugo- Villarino et al., 2010). While zebrafish do not have discrete lymph nodes, they do possess a lymphatic system with many characteristics of other vertebrate lymphatic systems (Yaniv et al., 2006). As previously mentioned, numerous lines are available with

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transgenic reporter genes for or knockouts of specific cell types, allowing for the visualization of specific cell types or the study of the contribution of specific immune cell populations to the host response to microsporidia in the zebrafish. One important and useful aspect of the zebrafish immune system is the temporal separation between the development of the innate and adaptive immune systems. The zebrafish embryo has been shown to have functionally competent macrophages as early as 1 day post fertilization (Herbomel et al., 2001), whereas while the adaptive immune system does not become fully functional until 4-6 weeks post fertilization (Lam et al., 2004). This temporal separation has been exploited in order to characterize differences in the zebrafish transcriptional response between innate and adaptive immunity to organisms such as Mycobacterium spp. (van der Sar et al., 2009). This characteristic of zebrafish could be similarly exploited to better understand the roles of innate and adaptive immunity in microsporidian infections. The pathological changes associated with microsporidian infections are highly dependent upon the immune status of the host. Whereas microsporidian infections of immune competent hosts generally begin with an acute phase which resolves, often

resulting in a long chronic phase, infections of immune compromised hosts are often

acute and lethal (Didier, 2000). A protocol has been developed whereby the immune

system is sub-lethally or lethally ablated using total body gamma irradiation (Taylor and Zon, 2009; Traver et al., 2004), allowing for the study of microsporidian infection in an experimentally immune-compromised host. Indeed, it was the use of total body irradiation which led to the discovery of Pleistophora hyphessobryconis in one zebrafish facility (Sanders et al., 2010), and this parasite is discussed in more detail below. Whereas this microsporidium rarely infects wild type zebrafish or results in chronic, subclinical infections, immune suppression by total body irradiation led to acute infection and dramatic mortality among these fish (Sanders et al., 2010). Interestingly, one isogenic line (CGL-1) also is highly susceptible to P. hyphessobryconis. The majority of studies around the host-pathogen interactions of microsporidian infections have examined acute infections and very little is currently

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known about factors which allow persistence of the parasite. The zebrafish infection model of P. hyphessobryconis appears to be amenable to the study of both acute and chronic infections as most infections are chronic, becoming acute after experimental immune suppression. Genomic/Transcriptomic The zebrafish is highly amenable to gene expression studies. Several studies have been performed to investigate the changes in gene expression of zebrafish in response to infection by pathogens. Whereas earlier studies of global gene expression made use of highly developed zebrafish microarrays made commercially or in-house (Pichler et al., 2004), more recent studies have made use of next-generation high throughput RNA- sequencing (RNA-Seq) technologies (Aanes et al., 2011; Hegedus et al., 2009; Ordas et al., 2011). These studies have provided a great deal of information regarding the transcriptional response of zebrafish to bacterial pathogens. The zebrafish genome assembly is currently in its ninth version and has incorporated transcript sequence data from numerous studies, providing a robust database from which to identify transcripts from RNA-Seq datasets. As the technology is maturing, RNA-Seq studies are able to

provide great insight into host-pathogen interactions on the level of host transcriptional

response. A recent study (Cuomo and Desjardins, 2012) examined the transcriptome of

host and pathogen, Caenorhabditis elegans and the microsporidum Nematocida parisii, respectively, during infection. A similar study using the zebrafish/Pseudoloma neurophilia model would provide information regarding the host and pathogen transcriptional response during infection in a vertebrate host. MICROSPORIDIAN INFECTIONS OF ZEBRAFISH There are currently two known species of Microsporidia that can naturally infect the zebrafish: Pseudoloma neurophilia and Pleistophora hyphessobryconis (Matthews et al., 2001; Sanders et al., 2010, 2012). These species are distantly related and, not surprisingly, produce profoundly different pathological presentations. Pseudoloma neurophilia is responsible for chronic infections of zebrafish and has a high prevalence in zebrafish research colonies. In contrast, Pleistophora hyphessobryconis is not

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encountered frequently in laboratory zebrafish facilities. Whereas it also generally results in chronic infections with little to no clinical signs, immune suppression can lead to very acute mortalities and specific fish lines appear to be more susceptible. Pseudoloma neurophilia The first report of a microsporidium infecting zebrafish was by de Kinkelin (de Kinkelin, 1980) in fish purchased from a pet store for use in a toxicological study over 30 years ago. Noting a high prevalence of spinal deformities such as lordosis and scoliosis and swimming abnormalities, de Kinkelin examined a number of these fish by histology and observed the presence of clusters of small, ellipsoidal spores, consistent with those produced by microsporidian parasites, localized to the ventral spinal cord. Subsequently, Pseudoloma neurophilia was described by Matthews et al (Matthews et al., 2001) in the ventral spinal cord, hindbrain, and skeletal muscle of both emaciated and healthy- appearing zebrafish housed at the Zebrafish International Resource Center (ZIRC). Since then, awareness of Pseudoloma neurophilia infections in laboratory zebrafish has grown dramatically. In 2011, infections of zebrafish by P. neurophilia were reported in 74% of zebrafish facilities examined through the ZIRC diagnostic service (Murray et al., 2011).

The concern of researchers for potential non-protocol induced variation (Kent et al.,

2012) in studies utilizing zebrafish with chronic microsporidian infection led to the

derivation of a zebrafish colony that is specific pathogen free (SPF) for P. neurophilia (Kent et al., 2011). In addition to providing SPF fish for toxicological research, this colony also provides the opportunity to study transmission of P. neurophilia using fish that are known to be P. neurophilia free prior to experimental exposure. As the name implies, Pseudoloma neurophilia, chronically infects neural tissue and develops into mature spores, forming aggregates mainly in the hindbrain, spinal cord, motor nerve ganglia and spinal nerve roots (Fig. 6.3). Zebrafish infected with P. neurophilia often have no visible signs of disease, however clinical signs such as emaciation and spinal deformity (Fig. 6.4) can occur. The parasite has been shown to enter the host via the intestinal epithelium and infects extraintestinal skeletal muscle myocytes in early stages of the infection. However, the host appears to be able to control

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the infection within myocytes as developing stages of P. neurophilia are generally found in immune-privileged sites such as the spinal cord and nerves. Presporogonic stages and mature spores are rarely found in skeletal muscle in later stages of the infection except in very acute, fulminant infections in which few parasites can be seen in histological sections to cause a severe myositis (Figs. 6.5, 6.6). However, the ultrastructure of P. neurophilia early developmental stages has been described within the sarcoplasm of somatic muscle cells in larval fish (Cali et al., 2012), and it has been described in many other organs including kidney and, most notably, ovarian stroma and within developing oocytes. So, while it can often be found in other tissue types, the apparent tropism of P. neurophilia for tissues found in immune-privileged sites such as the central nervous system and the eggs suggests that this evolved in response to pressure from the host immune system. P. neurophilia ultrastructure/development The development of Pseudoloma neurophilia typically begins by ingestion of an infectious spore. Spores of P. neurophilia are uninucleate and pyriform with an average length of 5.4 µm and width of 2.7 µm. They contain a large posterior vacuole and an

isofilar polar tubule with 14-17 coils arranged in a single row within the interior of the

spore (Cali et al., 2012). As with most microsporidian spores, these are extremely tough

structures meant to withstand long periods in the environment and possess a thick, chitinous endospore requiring special methods to disrupt them (Sanders and Kent, 2011). Cali et al. (Cali et al., 2012) described the sequential development of P. neurophilia based on experimental infections of zebrafish using electron microscopy. Once, ingested, the spores become closely associated with the villi of the intestine and presumably germinate, firing the polar tubule and injecting the sporoplasm into the cytoplasm of an intestinal epithelial cell. Proliferation of the sporoplasm occurs, in direct contact with the host cytoplasm beginning with several rounds of karyokinesis, producing a multinucleate plasmodial cell. Cytokinesis occurs in a stepwise fashion, resulting in several uninucleate cells which eventually undergo sporogony, indicated by the separation of the surface coat from the plasmalemmal surface. This surface coat forms the sporophorous vesicle

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(SPOV) in which another round of karyokinesis occurs, producing another multinucleated plasmodium, followed by cytokinesis, the formation of uninucleate sporoblasts and eventually the metamorphosis into spores. The SPOV is a fairly resilient structure which often maintains its integrity outside of the host tissue and is visible in wet mounts containing 8 to 16 spores (Fig. 6.7). The development of P. neurophilia appears to be relatively rapid with proliferative stages and sporonts observable at 4-5 days post exposure, and all stages, including mature spores, present at 8 days post exposure. We have recently developed an in situ probe for P. neurophilia based on the small subunit rDNA gene, allowing for the visualization of the distribution of presporogonic stages of the parasite in early infections. Experimental exposure of larval fish, followed by histological examination revealed that the parasite initially infects a wide variety of organs, including the intestine, kidney, pancreas and liver (Fig. 6.8). P. neurophilia Transmission As with many other microsporidia, the transmission of Pseudoloma neurophilia is more complex than originally thought (Fig. 6.9). Elucidation of the transmission characteristics of P. neurophilia has been important not only to the control of infections

in laboratory zebrafish colonies but also to the understanding of the basic biology of

microsporidian parasites. Our early studies showed that P. neurophilia is be transmitted

horizontally, ostensibly per os, by exposing adult zebrafish to inocula prepared from the spinal cords of infected fish (Kent and Bishop-Stewart, 2003). These results suggested that the cannibalism of dead tankmates serves to transmit the infection to naïve hosts. Later studies showed that infectious spores of P. neurophilia are also shed into the water by live fish during spawning (Sanders and Kent, 2011) and that fish cohabitating with infected fish become infected at a high rate (Sanders et al., 2012). Whereas P. neurophilia is clearly horizontally transmitted, the shedding of spores during spawning and the presence of spores in the ovarian tissue and within developing oocytes (Fig. 6.10) suggested the possiblity of vertical transmission of P. neurophilia (Kent and Bishop-Stewart, 2003; Sanders et al., 2012). Indeed, a more recent study (Sanders et al., in press), verified that that vertical transmission, both intraovum and

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extraovum, occurs. The low prevalence of intraovum transmission observed in single spawning events in this study (~1%) suggests that this mode of transmission plays only a minor role in the maintenance of P. neurophilia in a zebrafish population. However, it is important to note that zebrafish spawn continuously and the cumulative effect of even a low frequency of intraovum infections can result in a widespread infection, especially in populations where horizontal transmission is controlled. For example, laboratory fish colonies are often rigorously observed and fish exhibiting clinical signs of disease are removed, and most facilities introduce fish only as F1 generations from surface disinfected eggs (Harper and Lawrence, 2010; Kent et al., 2009; Lawrence, 2007). In addition, extraovum transmission (i.e., the shedding of spores during spawning), coupled with the high susceptibility of larval zebrafish, provides further opportunities for the parasite to be transmitted to novel hosts. Vertical transmission of pathogens in fish has serious implications for the control of a pathogen in aquaculture, but vertical transmission of P. neurophilia provides a vertebrate model in which to study evolutionary principles of virulence as they relate to mode of transmission. As P. neurophilia has been shown to be transmitted both

horizontally (by cohabitation and scavenging), and vertically (both intra- and extraovum),

studies in which the mode of transmission is experimentally manipulated could help to

elucidate important factors associated with virulence and mode of transmission of microsporidians in a vertebrate model system. The in vitro propagation of a pathogen is of obvious importance to the study of that pathogen. The long-term culture of microsporidia that infect insects and mammals has been achieved for numerous species. Unfortunately, in vitro culture of many fish- infecting species of the Microsporidia remains very challenging. Pseudoloma neurophilia has been grown on several fish cell lines, however, that growth is very limited (Monaghan et al., 2009) and relatively few spores are produced. Work is continuing on the development of an in vitro propagation system for P. neurophilia.

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Pleistophora hyphessobryconis In contrast to the high prevalence of P. neurophilia in laboratory zebrafish, Pleistophora hyphessobryconis is only rarely observed (Sanders et al., 2010). Commonly known as “neon tetra disease” for its type host, the neon tetra, Paracheirodon innesi, this myocyte-infecting microsporidium is widespread in the aquarium trade. P. hyphessobryconis infects a wide range of fishes in several families and has been reported from many species of aquarium fishes including Danio rerio (Sanders et al., 2010; Steffens, 1962). Infections of zebrafish by P. hyphessobryconis generally produce latent, subclinical infections at a low prevalence. The parasite infects myocytes and developmental stages can be seen throughout the skeletal muscle myofibers with focal areas of chronic inflammation (Fig. 6.11). These infections can become acute, resulting in prominent clinical signs and mortality, following incidental or experimental immunosuppression by gamma irradiation or in one particularly susceptible line, the CG1 isogenic line (Sanders et al., 2010). Clinical signs of acute P. hyphessobryconis infection include lethargy and the presence of focal depigmented regions, especially around the dorsal fin. These signs are rapidly followed by mortality and macroscopic examination

of the skeletal muscle can reveal large opaque areas consisting of necrotic myofibers

filled with P. hyphessobryconis spores (Fig. 6.12).

Transmission of P. hyphessobryconis is horizontal, ostensibly per os, and there is no evidence of intraovum transmission with this species. However, maternal transmission was suggested as infections were observed in 8 day-old neon tetras (Schaperclaus, 1991). Whereas the pathological changes caused by this microsporidium in zebrafish has been described (Sanders et al. 2010), little is known regarding the difference in tissue tropism between P. hyphessobryconis and P. neurophilia, the study of which would likely yield interesting results. Other microsporidia Numerous microsporidian species infect fishes (see Kent et al. Chapter 19), and many have a broad host range. Thus, there are likely many other species which can infect zebrafish, allowing for the study of a broad range of microsporidian associated diseases

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in a well-characterized vertebrate model. For example, recent work has shown that Anncaliia algerae, a microsporidium with an exceptionally broad host range including insects and human, can infect zebrafish cell lines (Monaghan et al., 2011). The ability for zebrafish to survive and remain immunologically competent within such a wide range of temperatures (Dios et al., 2010; Novoa et al., 2006) suggests that this model could be used to investigate other microsporidian species known to infect humans and other mammals. CONCLUSION While numerous microsporidia are known to infect mammals, many more have been recorded from fishes. The dramatic growth in the use of the zebrafish, Danio rerio, as a laboratory model for developmental, toxicological, cancer, and infectious disease research has increased interest in the pathogens of these fish. The most commonly diagnosed infectious disease of zebrafish is by the microsporidian parasite, Pseudoloma neurophilia, which is generally associated with chronic infections of laboratory zebrafish and has a high prevalence in zebrafish housed in research colonies. The presence of chronic infections in mammalian laboratory animals has been

identified as an incidental source of non-protocol induced variation and the same is true

for the zebrafish. The use of a well-developed vertebrate model to investigate host

response to microsporidian infection will allow the results from this study to provide insights into the host-pathogen interaction of microsporidian infections, which could be applied to microsporidian infections of other vertebrate animals. Microsporidians are not only important as a pathogen compromising zebrafish in research, but they also provide an excellent format for the study of microsporidia in a vertebrate model. For example, the zebrafish can be used for: 1) fine study of initial infection by intravital imaging, due to its small size and optical clarity, 2) high throughput in vivo screening of antimicrosporidial drugs, 3) the study of evolution of virulence as it relates to mode of transmission (vertical vs hortizontal) due to its high fecundity and rapid development, and 4) the genetics underlying susceptibility can be investigated at various “omic” levels using existing tools, with two very different types of

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parasites. The use of zebrafish as a model for the study of microsporidiosis is in its early stages and tools continue to be developed. The benefits of this vertebrate model system in the study of microsporidiosis are only just beginning to be realized and the future holds many exciting avenues of research using the zebrafish/microsporidian model system to discover numerous aspects of the Microsporidia.

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Figure 6.1. Histological section of a 7 day post-fertilization larval zebrafish. Numerous tissues can be visualized in this single section, including the liver (L), kidney (K), swim bladder (S), pancreas (K), eye (E) and gill (G). Hematoxylin and eosin stain, bar = 0.5 mm.

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Figure 6.2. Developing zebrafish embryo at 48 hours post-fertization with two opaque aggregates of Pseudoloma neurophilia (arrows). Bar = 1 mm.

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Figure 6.3. Histological section of an adult zebrafish showing aggregates of spores of P. neurophilia in the spinal cord (arrows). Hematoxylin and eosin stain, bar = 50 µm.

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Figure 6.4 Adult zebrafish, Danio rerio, infected with the microsporidian Pseudoloma neurophilia. While chronic infections with P. neurophilia can produce little to no grossly visible signs, clinical microsporidiosis can result in prominent emaciation and, occasionally, spinal deformation as can be observed. Bar = 0.5 cm.

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Figure 6.5. Histological section of an adult zebrafish with focal chronic inflammation and myodegeneration due to Pseudoloma neurophilia infection. Hematoxylin and eosin stain, bar = 40 µm.

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Figure 6.6. High magnification of a histological section of an adult zebrafish with chronic myositis caused by Pseudoloma neurophilia infection. Numerous single spores and small aggregates of spores (arrows) can be seen. Luna stain, bar = 40 µm.

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Figure 6.7. Wet mount of several aggregates of Pseudoloma neurophilia spores obtained from the hindbrain of an infected adult fish. Spores can be seen occurring within a sporophorous vesicle (arrows). Bar = 10 µm.

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Figure 6.8. Histological section of a larval zebrafish (7 days post fertilization) with developing stages of Pseudoloma neurophilia present in the liver visualized by in situ hybridization of a DIG-labeled single-stranded DNA probe which binds to a portion of

the P. neurophilia small-subunit ribosomal RNA gene. After binding, an anti-DIG alkaline phophatase conjugate is added which catalyzes a color reaction with 5-bromo-4 chloro-3-indoyl-phosphate (BCIP) and nitroblue tetrazolium salt (NBT) substrates,

producing a dark blue stain. Section is counterstained with Nuclear Fast Red (Vector Laboratories Inc, Burlingame, CA). h = hepatocyte nucleus, bar = 10 µm.

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Figure 6.9. Diagram illustrating the modes of transmission observed in Pseudoloma neurophilia infections in the zebrafish, Danio rerio.

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Figure 6.10. Histological section of an adult female zebrafish with mature spores (arrow) and presporogonic stages (p) of Pseudoloma neurophilia observed within a late-stage, vitellogenic oocyte. Luna stain, bar = 10 µm.

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Figure 6.11. Histological section of an adult zebrafish infected with Pleistophora hyphessobryconis. Focal inflammation can be observed (Inf). Hematoxylin and eosin stain, bar = 20 µm

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Figure 6.12. Adult zebrafish, Danio rerio, with skin removed infected with Pleistophora

hyphessobryconis. The flesh becomes opaque due to massive infection (arrow) and associated inflammation. Bar = 0.5 cm.

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Chapter 7. Summary and Conclusions The aims of this dissertation were to characterize the modes of transmission of Pseudoloma neurophilia in the zebrafish, Danio rerio and to describe the organ distribution of the parasite in early stages of infection in larval fish. As Pseudoloma neurophilia is the most commonly diagnosed infectious disease of laboratory zebrafish, the results of my studies are of great interest to the entire zebrafish research community and to the understanding of microsporidiosis in vertebrates. In developing the tools to achieve my research aims, namely a more sensitive real-time PCR based assay for the detection of P. neurophilia in fish tissues and in water, I have also provided the zebrafish community with a sensitive assay with which to test laboratory zebrafish for the presence of P. neurophilia. This assay has been used to develop and maintain the first P. neurophilia-free laboratory colony of zebrafish here at Oregon State University and is now in use in several laboratories worldwide to screen for this important pathogen. One of the key findings of my research is the verification of vertical transmission of P. neurophilia in the zebrafish and the observation of intraovum transmission. This is the first direct observation of vertical transmission of a microsporidian parasite in a

vertebrate, and has important implications both in the control of microsporidiosis in

laboratory zebrafish colonies and in the study of transmission and virulence of

microsporidia in vertebrates. The unequivocal presence of vertical transmission of P. neurophilia in zebrafish requires rigorous screening of brood fish for the parasite in order to establish a population free of P. neurophilia infection. Ultimately, the establishment of specific pathogen free fish colonies, such as the one here at Oregon State University, is the only reliable method for the exclusion of P. neurophilia from zebrafish used in research. The two modes of transmission of P. neurophilia in D. rerio: horizontal transmission by scavenging on dead fish and cohabitation with live fish, and the low, yet variable (0.2-1.5%), prevalence of intraovum vertical transmission as well as potential for extraovum transmission based on the shedding of high numbers of spores (~700-17 000) in several paired spawns, all indicate that this is a system which may be experimentally manipulated, allowing for studies of the evolution of virulence and its relatedness to

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mode of transmission in a well-described vertebrate animal model. These types of experiments have yielded interesting results using phage-bacteria and microsporidia arthropod model systems and now, with the characterization of modes of transmission in the zebrafish, this is possible in a vertebrate model which is widely used in biomedical research. Understanding the distribution and development of parasites within the host in the intial phases of infection is also important to understanding the biology and pathogenesis of these pathogens. Several microsporidia have relatively simply infection cycles whereby the entire process occurs within a single tissue such as the intestinal epithelium. This is true of certain species which infect humans such as Encephalitozoon intestinalis. Systemic microsporidiosis, however, occurs, especially among immunocompromised humans but also among many fishes. In zebrafish, latent P. neurophilia infections are especially problematic due to the potential for nonprotocol induced variation in studies using infected fish. My observation of P. neurophilia in tissues shortly after initial exposure provides an important clue as to the mechanism of infection. As I first observed sporulation at 4 d post exposure, it is unlikely that autoinfection occurs in the early stages

of P. neurophilia, rather the tissues in which the parasite is first observed are within the

range of the very long (over 150 µm) polar tubule from the intestine. The simultaneous

appearance of presporogonic proliferative stages of P. neurophilia in the liver, pancreas, and kidney indicates that these are the initial sites of infection, reached by the action of the polar tubule penetrating the wall of the intestine. The observation of parasite development within such a broad range of tissues also illustrates the potential for non- protocol induced variation presented by P. neurophilia infections in studies using larval zebrafish. Previous experiments tracking microsporidian progression in vertebrates have been limited by the size of the animal being studied, requiring the dissection of animals and examination of only partial organs. The use of larval zebrafish allowed me to examine several entire animals. The zebrafish is a rapidly growing model for bacterial, viral and fungal pathogens and its small size is one of its great strengths. The high

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prevalence of P. neurophilia in laboratory zebrafish makes it a logical biomedical model in which to study microsporidiosis. The results of my work characterizing the modes of transmission of Pseudoloma neurophilia in the zebrafish, and the organ distribution of the parasite in early phases further the development of the zebrafish as a model for microsporidiosis. The Microsporidia are a notoriously difficult group of organisms to study. As obligate intracellular pathogens, they have no development outside of an amenable host cell and, due to their unique method of transmission, relatively simple seeming procedures, such as the propagation of these parasites in cell culture, is not yet possible for the vast majority of microsporidian species. Even for the species which readily grow in cell culture, no known method of selective growth exists, making recombination and fluorescent tagging, something commonplace in numerous bacterial species today, impossible. Thus, microsporidian research must rely on relatively simple methods for the time being and the use of zebrafish as a vertebrate model for microsporidiosis makes this a much more tractable prospect.

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