Disease threats to Belgian anurans – Implications for amphibian conservation and human health

Mojdeh Sharifian Fard

Thesis submitted in fulfillment of requirements for the degree of Doctor in veterinary science (PhD), Faculty of Veterinary Medicine, Ghent University, 2014

Promoters: Prof. dr. An Martel Prof. dr. Frank Pasmans

Faculty of Veterinary Medicine Department of Pathology, Bacteriology and Poultry Diseases

Sharifian Fard Mojdeh Disease threats to Belgian anurans – Implications for amphibian conservation and human health 2014 Faculty of Veterinary Medicine, Ghent University Printed by University Press, Zelzate

ISBN: 9789058643889

Dedicated to my mother (Giti) and father (Hafez) In memory of Sepanta

CONTENTS

Table of Contents

INTRODUCTION ...... 1 1. Indigenous Amphibians of Belgium ...... 3 1.1. Indigenous species: definition ...... 3 1.2. Indigenous amphibians of Belgium ...... 3 1.2.1. Anura ...... 3 1.2.1.1. Alytes obstetricans, Common midwife toad ...... 4 1.2.1.2. Bombina variegata, Yellow-bellied toad ...... 5 1.2.1.3. Bufo bufo, Common toad ...... 6 1.2.1.4. Epidalea calamita, Natterjack toad ...... 7 1.2.1.5. Hyla arborea, Common tree frog ...... 8 1.2.1.6. Pelobates fuscus, Common spadefoot toad...... 9 1.2.1.7. Pelophylax lessonae, Pool frog ...... 10 1.2.1.8. Pelophylax kl. esculentus, Edible frog ...... 11 1.2.1.9. Rana arvalis, Moor frog ...... 12 1.2.1.10. Rana temporaria, Common frog ...... 13 1.2.2. Caudata ...... 14 1.2.2.1. Ichthyosaura alpestris, Alpine newt ...... 15 1.2.2.2. Lissotriton helveticus, Palmate newt ...... 16 1.2.2.3. Lissotriton vulgaris, Smooth newt ...... 17 1.2.2.4. Salamandra salamandra, Fire salamander ...... 18 1.2.2.5. Triturus cristatus, Great crested newt ...... 19 2. Invasive amphibians ...... 20 2.1. Invasive species: definition ...... 20 2.2. Invasive amphibians of Belgium ...... 21 2.2.1. Lithobates catesbeianus, American bullfrog ...... 22 2.2.2. Pelophylax ridibundus, Marsh frog ...... 24 3. Amphibian Diseases ...... 26 3.1. Ranavirosis ...... 27 3.1.1. Background ...... 27 3.1.2. General characteristics ...... 29 3.1.3. Replication cycle ...... 30 3.1.4. Transmission and pathogenesis ...... 31

I CONTENTS

3.1.5. Clinical signs and pathology ...... 33 3.1.6. Diagnosis ...... 35 3.1.7. Conservation strategies ...... 36 3.2. Chytridiomycosis ...... 37 3.2.1. Background ...... 37 3.2.2. Phylogenetic characteristics of B. dendrobatidis ...... 40 3.2.3. Lifecycle stages ...... 40 3.2.4. Susceptibility to B. dendrobatidis ...... 41 3.2.5. Clinical signs, gross pathology and pathogenesis ...... 43 3.2.6. Transmission ...... 44 3.2.7. Amphibian defense against chytridiomycosis ...... 44 3.2.8. Diagnosis ...... 45 3.2.9. Preventative management strategies ...... 46 3.3. Zoonotic bacterial diseases ...... 48 3.3.1. Chlamydia ...... 48 3.3.2. Salmonella ...... 49 3.4. Mesomycetozoans ...... 50 3.5. Parasites ...... 50 3.6. Non-infectious diseases ...... 53 3.6.1. Heavy metal disorders ...... 53 3.6.1.1. Trace metals ...... 53 3.6.1.1.1. Copper ...... 54 3.6.1.1.2. Zinc ...... 55 3.6.1.2. Xenobiotic metals ...... 57 3.6.1.2.1. Lead ...... 57 3.6.1.2.2. Mercury ...... 58 3.6.1.2.3. Cadmium ...... 59 4. References ...... 61

SCIENTIFIC AIMS ...... 89

STUDY 1 ...... 93

Road-killed common toads (Bufo bufo) in Flanders (Belgium) reveal low prevalence of ranaviruses and Batrachochytrium dendrobatidis

II CONTENTS

STUDY 2 ...... 103

Ranavirosis in invasive bullfrogs, Belgium

STUDY 3 ...... 109

The novel ‘Candidatus Amphibiichlamydia ranarum’ is highly prevalent in invasive exotic bullfrogs (Lithobates catesbeianus)

STUDY 4 ...... 119

Chironomidae bloodworms larvae as aquatic amphibian food

STUDY 5 ...... 139

Migrating common toads (Bufo bufo) in rural temperate regions: reservoirs of Salmonella?

STUDY 6 ...... 151

The absence of zoonotic agents in invasive bullfrogs (Lithobates catesbeianus) in Belgium and The Netherlands

GENERAL DISCUSSION ...... 163

SUMMARY ...... 177

SAMENVATTING ...... 181

CURRICULUM VITAE ...... 187

BIBLIOGRAPHY ...... 191

ACKNOWLEDGEMENTS ...... 195

III

LIST OF ABBREVIATIONS

ADF acid detergent fibre AMP anti-microbial peptide ANOVA analysis of variance ATV Ambystoma tigrinum virus Bd Batrachochytrium dendrobatidis Bs Batrachochytrium salamandrivorans BdCAPE Batrachochytrium dendrobatidis Cape lineage BdCH Batrachochytrium dendrobatidis Swiss lineage (BdCH) BdGPL Batrachochytrium dendrobatidis Global Panzootic lineage BIV Bohle iridovirus Ca calcium Cd cadmium CD8 cluster of differentiation 8 CF crude fat CMTV common midwife toad virus COI mitochondrial cytochrome oxidase subunit CP crude protein CTOH chytrid-thermal-optimum hypothesis Cu copper 2,4-DAPG 2,4-diacetylphloroglucinol DE delayed early delta-ALAD delta-aminolevulinic acid dehydratase DNA deoxyribonucleic acid DM ethylenediaminetetraacetic acid EDTA dry matter ELISA enzyme-linked immunosorbent assay EM electron microscopy EPH endemic pathogen hypothesis FEP free erythrocyte protoporphyrin FV3 Frog virus 3 GC guanine-cytosine GIV Grouper iridovirus Hg mercury HPLC high performance liquid chromatography HS haemorrhage syndrome I3C indol-3-carboxaldehyde IE immediate early IG immune Globulin IHC immunohistochemistry ISEIA invasive alien species in Belgium IUCN international union for conservation of nature L viral transcripts late viral transcripts MCP major capsid protein MeHg methylmercury mRNA messenger ribonucleic acid MT metallothionein NCBI national center for biotechnology information NDF neutral detergent fibre NH4 ammonium NOX nitrogen oxide NPH novel pathogen hypothesis

IV LIST OF ABBREVIATIONS

OIE world organisation for animal health ORF open reading frames PAS periodic acid-Schiff Pb lead PCR polymerase chain reaction PO per oral Q-PCR quantitative PCR Q–Q plot quantile-quantile plot RCRZ Rana catesbeiana virus Z RNA ribonucleic acid RNA Pol II RNA polymerase II ROS reactive oxygen species SGIV Singapor grouper iridovirus SH sulphur SO2 sulphur dioxide SSU-rDNA small subunit ribosomal DNA TAE tris-acetate TLR toll-like receptor Uk United kingdom US ulcerative syndrome UV ultra violet ZIP zinc-iron-related transporter proteins Zn zinc

V LIST OF ABBREVIATIONS

VI

INTRODUCTION

INTRODUCTION

2 331 INTRODUCTION

1. Indigenous Amphibians of Belgium 1.1. Indigenous species: definition Indigenous means originating and living or occurring naturally in an area or environment. A species is defined as indigenous to a given region or ecosystem if its presence is the result of only natural processes, without human interference (Drake and ICSU, 1989; Cox, 1999).

1.2. Indigenous amphibians of Belgium The class Amphibia (Amphi: on both sides, bios: life) currently consists of 6,915 species divided in three orders: the Anura (frogs and toads) (5,602 species, 48 families) (88%), the Caudata or Urodela (salamanders and newts) (571 species, 10 families) (9%), and the Gymnophiona or Apoda (caecilians) (174 species, 3 families) (3%) (Frost, 2013). The fourth order, the Temnospondyli consists of primitive amphibians from the Carboniferous, Permian and Triassic periods and is completely extinct (Palmer, 1999). Belgium hosts 15 species of amphibians, including 10 species of Anura (5 toads and 5 frogs) and 5 Caudata species (4 newts and 1 salamander) (Tables 1 and 2) (Vences, 2004; Frost, 2013). All known amphibian species have been assessed using the International Union for Conservation of Nature (IUCN) Red List categories and criteria. These categories provide an explicit framework for determining a species' conservation status, with an emphasis on identifying those at highest risk of global extinction. In this context, the term "Threatened" refers to those species classified under IUCN Red List categories of Vulnerable, Endangered, or Critically Endangered. In Belgium, Red Lists are compiled by the Research Institute for Nature and Forest according to IUCN criteria. It noted that among the 15 indigenous species present, 10 of these are decreasing in population and are currently threatened. The last assessment in 2004 indicated that 38% of amphibian species are considered to be threatened species in Belgium, which can be broken down regionally as 25% in Flanders, 27% in Wallonia and 40% in Brussels (Kestemont, 2010).

1.2.1. Anura In the Anura (meaning “tail-less”, from the Greek an-, without + oura, tail), a distinction is made between frogs and toads on the basis of their appearance, but this has no

3 INTRODUCTION taxonomic basis (Frost, 2013). There are 10 indigenous species of Anura in Belgium (Kuijken et al., 2001). Table 1 gives an overview of the status of indigenous Anura in Belgium.

Table 1. Indigenous Anura and their status in Belgium Scientific name Common name Family Authority Status

Alytes obstetricans Common midwife toad Alytidae Linnaeus, 1758 Endangered

Bombina variegata Yellow-bellied toad Bombinatoridae Linnaeus, 1758 Critically endangered

Bufo bufo Common toad Bufonidae Laurenti, 1768 Safe/Low Risk

Epidalea calamita Natterjack toad Bufonidae Laurenti, 1768 Rare

Hyla arborea Common tree frog Hylidae Linnaeus, 1758 Critically endangered

Pelobates fuscus Common spadefoot toad Pelobatidae Laurenti, 1768 Critically endangered

Pelophylax lessonae Pool frog Ranidae Camerano, 1882 Endangered

Pelophylax kl. esculentus Edible frog Ranidae Linnaeus, 1758 Safe/Low Risk

Rana arvalis Moor frog Ranidae Nilsson, 1842 Rare

Rana temporaria Common frog Ranidae Linnaeus, 1758 Safe/Low Risk

1.2.1.1. Alytes obstetricans, Common midwife toad Alytes obstetricans (Figure 1A) gets its name from Greek and Latin. Alytes means “linked” or “entwined” in Greek; this name was given to them because the eggs are linked together and then entwined around the male’s hind legs. Obstetricans is Latin for “midwife”; the reason for this name is that the male inserts one of his toes into the female’s cloaca in order to help her lay the eggs. The common midwife toad can be distinguished by its large head and small, stubby, compact body with rough, reddish, warty skin (Nöllert and Nöllert, 1992). Reddish warts often extend from the tympanum to the groin (Arnold, 2003). The ventral colouration is dirty white, while the throat and the chest are often dappled with grey. The eyes have a vertical, slit-shaped pupil. The parotoid glands are less obvious than in other toads, but they are present at the top of the head in some specimens. The tympanum is mostly visible to the naked eye (Kwet, 2009). Although the appearance of the male and female is roughly similar, except during the breeding season, they can be differentiated by measuring three variables: the distance between nostrils, the distance between the anterior end of the middle metacarpal tubercle and the tip of the third finger, and the distance from the elbow to the third fingertip (Bosch and Marquez, 1996; Gasc et al., 1997). In the breeding season from March until August, males can be easily characterised by their unusual reproduction behaviour: they carry the eggs themselves (Nöllert

4 INTRODUCTION and Nöllert, 1992). Within its native range, Alytes obstetricans is strictly terrestrial and can live in temperate forests, semi-arid areas, walls, embankments, and slopes with small stones and sparse vegetation (Spellerberg, 2002; Gasc et al., 1997). Copulation and the fertilisation of the eggs are done on land and the male carriers the eggs around the hind legs. Approximately 3-6 weeks after mating, the male will begin to look for aquatic environments to deposit the tadpoles when they hatch (Gasc et al., 1997; Arnold, 2003). The common midwife toad is nocturnal by nature. Hibernation starts in October and lasts until late March. Mating activity often occurs in May and June (Spellerberg, 2002). The common midwife toad is currently threatened in Belgium and is listed as an endangered species (Kestemont, 2010). In Wallonia, the species has an ample distribution and populations are generally large. However, populations close to the Wallon-Flemish border seem to be in some form of decline. In Flanders, the midwife toad is restricted to several small populations at the periphery of it distributional range in Wallonia (Louette and Bauwens, 2013).

1.2.1.2. Bombina variegata, Yellow-bellied toad Bombina variegata (Figure 1B) is characterised by a vividly yellow belly with large, dark spots and by the bright, inner surface of the legs where they merge with the thigh (Gasc et al., 1997; Arnold, 2003). Bombina originates from the Latin for “hum”, indicating their voice. Variegata means “varied”, “different”, or “irregular” and was given to them because of the asymmetrical figure on their belly. These colour characteristics are the most important identification keys. The tips of the toes are covered with a brilliant yellow colour. The dorsal appearance is yellowish to brownish, with small dark points. They have a flat shape with warty dorsal skin and legs. The pupil has a triangle shape. (Nöllert and Nöllert, 1992; Gasc et al., 1997). The tympanic membrane and parotoid glands are absent. During the breeding season, dark nuptial pads appear on the males’ first, second and third toes (Spellerberg, 2002). Bombina variegata is found in terrestrial and aquatic habitats. They mainly inhabit low hills or lower altitudes in mountains with different types of bodies of water (e.g. lakes, ponds, swamps, rivers and streams, pools and springs) (Gasc et al., 1997). The pattern of spawn deposition can be interpreted as an attempt to select sites that allow for rapid larval development (warm water) and to avoid sites with high numbers of newts and invertebrate predators (Barandun and Reyer, 1997). Obligate hibernation starts between September and October and can last until May. Breeding begins 5-10 days after entering the aquatic

5

INTRODUCTION environment and it can last until August (Gasc et al., 1997). Bombina variegata uses various ponds for reproduction; however, it is described as a temporary pond breeder (Hartel, 2008). Bombina variegata is listed as a critically endangered species in Belgium (Kestemont, 2010). In Flanders, this species was historically restricted to the extreme eastern part, where it was last observed in the early 1980s (regionally extinct). In Wallonia, a remarkable decline in the size and number of its populations was observed as early as about 1880. This decline continued throughout the 20th century and at presence, only three very small populations survive (Louette and Bauwens, 2013).

1.2.1.3. Bufo bufo, Common toad Bufo bufo (Figure 1C) (Bufo is Latin word for “toad”) is a large genus of so-called “true toads” and is characterised by warty skin that is extensively covered by rounded tubercles on the surface. In particular, they possess warty glands behind the eyes (Gasc et al., 1997; Arnold, 2003). The common toad has a large, chubby body structure. Males have a nuptial pad on the first finger (Nöllert and Nöllert, 1992). These pads become visible on the second and/or third toes behind the front limbs throughout the breeding season. Females have no nuptial pads. If a male is picked up, he will start to squeak; however, the female does not react. The oblique parotoid glands, located behind the eyes, are visible. The tympanic membrane, however, is barely noticeable. The eyes often have a reddish-brown iris with a horizontal pupil (Vitt and Caldwell, 2008). There are two subarticular tubercles on the second and third toes of the fore feet. The tarsal fold is absent. Depending on their habitat, skin colouration can vary from brown/dark brown to grey with more or less developed dark blotches extending to the flanks. Sometimes, they are reddish in some regions (particularly if iron oxide exists in the soil) and even yellowish, often with many small dark spots along the dorsum. The underside is usually whitish or pale grey and often has a marbled effect. The colouration of body often changes to a single colour during the breeding season (Nöllert and Nöllert, 1992; Gasc et al., 1997). Bufo bufo is a widespread species and well-adapted to different habitats, even one that is very urbanised. Its habitat preferences are open woodlands (coniferous, mixed and deciduous forests) and/or rough grasslands (Spellerberg, 2002). In breeding seasons, they move to an aquatic habitat, including rivers and streams with relatively clear water in which to lay the eggs (Nöllert and Nöllert, 1992; Arnold, 2003). Hibernation occurs singly or in groups. Depending on the elevation, they hibernate from September/October to March/June (Spellerberg, 2002). However, some specimens are active

6 INTRODUCTION from late February onward. They hibernate in holes in woodlands, such as log piles, compost heaps and stone walls. Depending on local weather conditions, reproduction takes place from February (Nöllert and Nöllert, 1992). There are currently no major risk threats to Bufo bufo in Belgium and it is listed as being a safe/low risk species in both Flanders and Wallonia, although it is considered a vulnerable species in Brussels (Kestemont, 2010).

1.2.1.4. Epidalea calamita, Natterjack toad Epidalea calamita, formerly Bufo calamita (Figure 1D) is a typical toad that is characterised by a thin, yellow dorsal stripe running down the middle of the back (Nöllert and Nöllert, 1992; Arnold, 2003). Epidalea is of unknown origin. Calamita is Latin and means “reed”, referring to its habitat. The Natterjack gets its common name from its very loud and distinctive mating call during the breeding season, which is an attempt to attract female mates in the evening (Beebee, 1983; Vitt and Caldwell, 2008). It is described as one of Europe’s noisiest amphibian since its chorus can be heard over several kilometres. The limbs are relatively short, enabling them to run rather than walk, which is why they are sometimes referred to as the “running toad” (Arnold, 2003). The skin bears numerous irregular warts varying in colour from red, dark brown or green. The natterjack toad has golden-green eyes with horizontal pupils (Nöllert and Nöllert, 1992). Parallel parotoid glands are noticeable at the back of the eyes. The tympanic membrane is visible. General dorsal colouration is dark green or brown, while the warts are lighter in colour. The belly is whitish with small, dark green spots (Beebee, 1983). Males have stronger forelimbs than females. In breeding season, the males have dark nuptial pads on three fingers (Beebee, 1983). Epidalea calamita prefers to live in warm, open and unshaded areas with soft ground and low vegetation. Their aquatic habitats consist of small, shallow, temporary pools or even slightly brackish waters. It is typically found in lowland heaths close to shallow water where there are few predators such as fish or invertebrates (Beebee, 1983; Nöllert and Nöllert, 1992). Depending on the latitude of the habitat, breeding season occurs in April/July (Spellerberg, 2002). In Flanders, this specis is restricted to the western part of the coastline, anthropogenically elevated terrains near the harbor of Antwerp, and the main heathland areas in the Campine ecoregion. Over the years, the extensive landscape of Flanders changed in favour of grey infrastructure in combination with an intensified agricultural activity. In

7

INTRODUCTION

Wallonia, the species was once a common member of the amphibian fauna but now only occurs in five distinct clusters and is considered as endangered species (Louette and Bauwens, 2013).

1.2.1.5. Hyla arborea, Common tree frog Hyla arborea (Figure 1E) can be recognised by its bright green colour with dark lines extending down the flanks (Gasc et al., 1997; Arnold, 2003). Hyla originates from the Greek meaning “to bark” because this frog’s call is similar to a bark. Arborea is Latin for “tree” or “of the trees”. They are tiny and slim frogs, with especially thin hind limbs and very smooth skin. Depending on the substrate colour and temperature, the skin colour is usually lime green with dorsolateral stripes, which are long, thin, dark bands with a white outer edging running from the nostril towards the groin. This band usually forms an inguinal loop (Nöllert and Nöllert, 1992; Kuzmin, 1995). Some specimens have small spots, which are darker than the skin colour, located along the dorsal area. Although colouration of the majority of specimens is bright green, the skin colour can be darker in winter due to their need to heat up faster in the colder climate. Their eyes are golden/yellow with a horizontal pupil. The ventral surface is yellowish or whitish with a granular pattern (Tarkhnishvili and Gokhelashvili, 1999; Arnold, 2003). Males can be distinguished from females by a large, yellowish, guttural vocal sac, which is externally visible even when not in use (Nöllert and Nöllert, 1992). Well-developed adhesive discs for climbing cover the tips of every toe. Vomerine teeth and tympanic membranes are present. They have no parotoid glands or dorsolateral folds (Gasc et al., 1997). As the name indicates, the tree frog inhabits an arboreal environment. They spend a major part of their lives in broad-leafed and mixed forests with tall trees near streams and lake shores where the average temperature is around 12°C. They can also live in low vegetation (shrubs) (Spellerberg, 2002). They leave the vegetation and come down to the ground to forage, rehydrate and to hibernate. Hibernation usually starts in September/December and lasts until February/early May on land, e.g. in burrows, while spawning occurs in water such as ponds, ditches and puddles. The breeding season occurs between April and June when they come down from the top of trees to the ground (Terent'ev and Chernov, 1965; Spellerberg, 2002). The tree frog has been disappearing from its distribution range in Belgium over the last few decades. Although this species is considered as extirpated in Wallonia, as the species is

8 INTRODUCTION still present in Luxemburg, near the frontier of Wallonia, collaboration with this country was recently established to extend the existing populations. In Flanders, where formerly numerous populations existed, only few of them survived (Louette and Bauwens, 2013).

1.2.1.6. Pelobates fuscus, Common spadefoot toad Pelobates fuscus (Figure 1F) is commonly known as the common spadefoot toad or garlic toad (Arnold, 2003). Pelobates in Greek is composed of two words, the first meaning 'mud' and the second meaning to 'walk'. Fuscus comes from the Latin for “brown”. This species is characterised by a bulky body structure, a large head and reddish spots along the flanks (Nöllert and Nöllert, 1992; Gasc et al., 1997). Although the skin colouration is dependent on their habitat and gender, dorsal colouration usually varies between light grey and brown/beige, which is mottled by large, dark brown stripes with small, reddish points (Arnold, 2003). Moreover, the dorsal pattern is extremely variable between specimens, but usually the blotches along the body are dark (dark brown or grey) and the interval between blotches is often very light (yellowish, cream, olive, light brown or greenish). The belly is white or white-greyish with a mottled pattern (Nöllert and Nöllert, 1992; Gasc et al., 1997). The male dorsal colouration shows a tendency towards a yellowish colour, while females are more greyish. In contrast to most toads, their skin is smooth. However, a few distinct tubercles have been seen occasionally (Gasc et al., 1997). They have big, golden eyes with vertical pupils. The parotoid glands, tympanic membrane and vocal sac are absent. The extensive webbing between the toes of the hind feet is present (Arnold, 2003). Spadefoot toads get their name from the blade-like shape of the lower hind limbs, which have a nail-like formation that looks similar to a blade or spade. The colour of this part is often light cream (Nöllert and Nöllert, 1992; Gasc et al., 1997). During the breeding season, the males possess prominent oval glands and small tubercles on the forelimbs (Nöllert and Nöllert, 1992). Pelobates fuscus have a preference for open, often sandy or loose soiled ground where they can dig easier (Spellerberg, 2002). Hibernation starts in September/October and lasts until March. The breeding season lasts from March until April, although it can extend to June in some specimens. Reproduction often occurs around ponds, water holes, puddles and lakes (Nöllert and Nöllert, 1992; Arnold, 2003). Pelobates fuscus is a critically endangered species in Belgium. No observations have been recorded since 1987 in Wallonia and the population is now considered to have been extirpated. In Flanders, the presence of the spadefoot toad could be confirmed at only for

9 INTRODUCTION localities (Louette and Bauwens, 2013).

1.2.1.7. Pelophylax lessonae, Pool frog Pelophylax lessonae (Figure 1H) is a medium-sized, European frog (Arnold, 2003). Pelophylax originates from Greek and is composed of two words: the first being “mud” and the second “guardian”. This name was given to these frogs because they never move far away from the water and they often stay immobile on the water’s edge. Lessonae refers Michele Lessona, an Italian herpetologist. In comparison with other Pelophylax species, the pool frog is brighter in colour, making for a more vivid appearance. The skin colouration and patterns vary considerably from marbled brown to brownish-green with dark blotches. The dorsal colouration can be yellowish-green, olive green or greyish-green in some specimens. The ventral surface is white or yellowish-white, usually without blotches. Both the dorsolateral folds and middorsal line are visible (Nöllert and Nöllert, 1992; Gasc et al., 1997). Pool frogs have a sharp snout and a light stripe running down the middorsal surface. The vomerine teeth, ossified sternum, webbed toes and golden eyes with horizontal pupils are obvious. The posterior part of the tongue is free and forked. The heel of the short hind limb does not reach beyond the eye when stretched. Although the pool frog has webbed feet, their hind feet are not as well developed as other Pelophylax species. They lack temporal markings and parotoid glands. Pool frogs are diurnal by nature (Nöllert and Nöllert, 1992; Arnold, 2003). Males have nuptial pads on the first finger, which are greyish in colour and become darker during the breeding season. Moreover, males have a paired, whitish external vocal sac on either side of the mouth. Although both sexes become brighter and more yellowish during breeding season, the males are more vivid or even extremely yellow (Nöllert and Nöllert, 1992; Arnold, 2003). The pool frog is commonly mistaken for the edible frog, Pelophylax kl. esculentus, even though the pool frog has a larger inner metatarsal tubercle and longer legs that reach to the eyes and nostrils (Nöllert and Nöllert, 1992; Gasc et al., 1997). Pelophylax lessonae generally inhabits stagnant bodies of water, such as lakes, sunny ponds, swamps, large puddles and ditches close to vegetated areas. They can be found in or around the water throughout the year. The presence of permanent water is necessary for the existence of Pelophylax lessonae populations (Spellerberg, 2002). Pool frogs hibernate on land, usually in frost-free holes, from October to April. They can move from one hibernation site to another during the winter. Sometimes they can move over 100-200 meters. Reproduction extends from late March/May to late June. Mating occurs in aquatic sites such as water holes, ponds, lakes and other bodies of water (Gasc et al., 1997; Arnold, 2003).

10 INTRODUCTION

Pelophylax lessonae is listed as an endangered species in Belgium, particularly in Wallonia. It is a rare species in Flanders (Kestemont, 2010).

1.2.1.8. Pelophylax kl. esculentus, Edible frog Pelophylax kl. esculentus (Figure 1G) is the scientific name of the common European frog (Nöllert and Nöllert, 1992). Pelophylax: refers to Pelophylax lessonae. kl. is the abbreviation for “klepton” and comes from the Latin “to steal” or “to conceal”, referring to the chromosomes that are “stolen” in order to create a different frog when Pelophylax ridibundus and Pelophylax lessonae interbreed. Esculentus is Latin and means “edible”. This species is also known as the common water frog or green frog (Gasc et al., 1997; Arnold, 2003). Pelophylax kl. esculentus is produced as a result of interbreeding between Pelophylax lessonae (Pool frog) and Pelophylax ridibundus (Marsh frog) (Berger, 1988). However, the offspring of edible frogs are often malformed (Berger, 1988). Morphological traits of this hybrid are intermediate between the parent species, Pelophylax lessonae and Pelophylax ridibundus (Nöllert and Nöllert, 1992; Gasc et al., 1997). The dorsal colouration is usually marbled green with brown spots. They often have a light stripe running along the middorsal surface. The edible frog has golden eyes, a sharp snout, light grey vocal sacs on both sides of the mouth, and a white or light grey belly with dark spots (Tarkhnishvili and Gokhelashvili, 1999). The posterior heels can reach between the eyes and nostrils when the hind leg is stretched along the body. The inner metatarsal tubercle is nearly half the length of the first toe (1.73-2.89 times shorter than the first toe) (Arnold, 2003). Pelophylax kl. esculentus is a complex hybridogenetic form including diploid and triploid forms. Triploid forms display closer traits to either the Pelophylax lessonae or Pelophylax ridibundus from which they inherit an extra set of chromosomes (Kierzkowski et al., 2011). Males can be sexually distinguished from females due to the fact that the males have grey nuptial pads on the interior limbs, a pair of vocal sacs on both sides of the mouth, robust limbs with a small body, and well-developed metatarsal tubercles (Arnold, 2003). In addition, the male’s dorsal skin becomes extremely green in colour during breeding season (Nöllert and Nöllert, 1992; Arnold, 2003). Ecological parameters related to the edible frog are intermediate between the two parent species (Kuzmin, 1995). In general, they can inhabit all types of aquatic bodies where pool frogs and marsh frogs could be found, such as overgrown ponds, lakes, streams and channels sheltered by enough vegetation in a sunny location (Gasc et al., 1997; Arnold, 2003). They usually inhabit mixed populations with either pool frogs or marsh frogs, but they rarely live

11 INTRODUCTION with both of these species. The edible frog is diurnal and, in contrast to other water frogs, they can move far from water for hunting and feeding, but they do not enter dense forests (Spellerberg, 2002). Hibernation starts in September/October and can last up to March/May (Gasc et al., 1997). The place of hibernation is variable between the specimens. When Pelophylax kl. esculentus coexists with Pelophylax lessonae, hibernation occurs on land, whereas it can occur in water if they coexist with Pelophylax ridibundus. Mating calls in Pelophylax kl. esculentus are more similar to Pelophylax lessonae than Pelophylax ridibundus (Nöllert and Nöllert, 1992; Arnold, 2003). There are currently no major risk threats to Pelophylax kl. esculentus in Belgium. The conservation status of this species is categorised as lower risk (Bauwens and Claus, 1996). The edible frog has been protected in Flanders since 1980, and is listed as rare in Wallonia and regionally extinct in Brussels (Kestemont, 2010).

1.2.1.9. Rana arvalis, Moor frog Rana arvalis (Figure 1I) is characterised by a dark, temporal “bandit-like” band and a chevron-shaped (^) spot on the neck. Rana is Latin for “frog”. Arvalis is Latin and means “of the fields”. Its skin is smooth with a reddish-brown, grey, light olive, yellowish or rufous colour and prominent dorsolateral folds (Nöllert and Nöllert, 1992). There can be a pale stripe down the centre of the back (Arnold, 2003). The skin can be spotted with dark, irregular markings on the dorsolateral surface that vary in number, arrangement and size. The ventral surface is usually unspotted and white or yellow in colour. However, some specimens have an irregular pattern running along the belly (Gasc et al., 1997). Rana arvalis has short hind limbs and its head is more tapered than of Rana temporaria. The posterior part of the tongue is forked and free. The vomerine teeth and horizontal pupils are noticeable. The parotoid glands are absent. The hind feet are webbed (Gasc et al., 1997; Arnold, 2003). Males can be sexually distinguished from females due to the nuptial pads (greyish in colour) on the first finger and a small pair of internal vocal sacs. The males’ skin colour changes to blue during breeding season (Nöllert and Nöllert, 1992). The moor frog inhabits open areas, including forest tundra, forest steppe edges and even semi-deserts (Spellerberg, 2002). They exhibit wide ecological plasticity and are more recognised as a thermophilous species than the sympatric common frog. Rana arvalis is considered to be a typical lowland species, reaching a maximum altitude of between 800 and 900 m (Gasc et al., 1997). Depending on the latitude of the location, hibernation occurs from

12 INTRODUCTION

September/November to February/March. Reproduction occurs between March and June in shallow, warm sites (Arnold, 2003). Although the moor frog has been a protected species since 1980, they are rare in Belgium now (Bauwens and Claus, 1996; Kestemont, 2010).

1.2.1.10. Rana temporaria, Common frog Rana temporaria (Figure 1J) is known as the European common frog or European common brown frog (Arnold, 2003). Rana: refers to Rana arvalis. Temporaria is Latin and means “temporary”; this name was given to them due to the large numbers of this species during the breeding season, after which it seems that they retreat and live solitary lives the rest of the time. The most distinguishing features of Rana temporaria include smooth, brownish skin with a dark marking running from the snout through the tympanum and a blunt, short snout (Nöllert and Nöllert, 1992; Arnold, 2003). Dorsal colouration can vary among some specimens, ranging from brownish, yellowish and even bluish with small dark blotches. The hind limbs often have a granular, brownish appearance. The body structure is corpulent (Gasc et al., 1997). Rana temporaria has vomerine teeth, webbed hind feet, ossified sternum, discrete dorsolateral folds, golden eyes with horizontal pupils, and a long first finger. The posterior section of the tongue is free and forked. The ventral surface is usually white or grey with a brownish-grey or almost black blotched pattern. The tibio-tarsal articulation on the hind limb commonly reaches the eye. The common frog has no parotoid glands and the light middorsal strip is usually absent (Kuzmin, 1995; Arnold, 2003). Males can be distinguished from females because the males have specific characteristics, including a pair of internal vocal sacs, more robust anterior limbs and dark brown/black nuptial pad on the first finger. During the breeding season, the males’ throats turn a bluish colour. The dorsal colouration often changes and becomes light and greyish, while females are more brownish or reddish (Nöllert and Nöllert, 1992; Gasc et al., 1997; Arnold, 2003). Rana temporaria inhabits both lowland and upland regions, such as deciduous or mixed forests, true tundra near permanent lakes, dry and swampy meadows, marshlands and moorlands (Spellerberg, 2002). They are found up to 3,000 m in altitude and prefer cooler areas rather than humid regions. If they inhabit a forested zone, they prefer the dense parts of it and behave as a hygrophilous species. Common frogs can also live in agricultural areas and even cities (Nöllert and Nöllert, 1992; Spellerberg, 2002). Depending on the latitude

13 INTRODUCTION and altitude, hibernation starts between October/early November, lasts up to February/early April, and occurs under the water in groups. Reproduction occurs from March - late June (Kuzmin, 1995). There are currently no major risk threats to Rana temporaria in Belgium. The conservation status of this species is categorised as lower risk and their populations are sufficiently high to ensure its survival in the long run (Bauwens and Claus, 1996). The common frog has been protected in Flanders since 1980, while it is listed as a vulnerable species in Brussels and as near-threatened in Wallonia (Kestemont, 2010).

Figure 1. Native Anura: Alytes obstetricans (A; credited by Frank Pasmans), Bombina variegata (B; credited by Frank Pasmans), Bufo bufo (C; credited by Frank Pasmans), Epidalea calamita (D; credited by Frank Pasmans), Hyla arborea (E; credited by Frank Pasmans), Pelobates fuscus (F; credited by Frank Pasmans), Pelophylax lessonae (G; credited by Andreas Nollert), Pelophylax kl.esculentus (H; credited by Frank Pasmans), Rana arvalis (I; credited by Maciej Bonk), Rana temporaria (J; credited by Frank Pasmans).

1.2.2. Caudata Caudata (Latin for “tail”) the second major extant order of the class Amphibia, with 571 species that are divided taxonomically into 10 families based on phylogeny and anatomical features. It includes salamanders and newts (Frost, 2013). Belgium hosts five species of Caudata, i.e. four newts and one salamander (Table 2) (Kuijken et al., 2001).

14 INTRODUCTION

Table 2. Indigenous Caudata and the current status in Belgium Scientific name Common name Family Authority Status

Ichthyosaura alpestris Alpine newt Salamandridae Laurenti, 1768 Safe/Low Risk Lissotriton helveticus Palmate newt Salamandridae Razoumowsky, 1789 Near-threatened Lissotriton vulgaris Smooth newt Salamandridae Linnaeus, 1758 Safe/Low Risk Salamandra salamandra Fire salamander Salamandridae Linnaeus, 1758 Vulnerable Triturus cristatus Great crested newt Salamandridae Laurenti, 1768 Rare

1.2.2.1. Ichthyosaura alpestris, Alpine newt Ichthyosaura alpestris (Figure 2A), formerly Triturus alpestris (Ichthyosaura originates from Greek and is composed of two words: Ichthyos means “fish” and Sauros means “lizard”. Alpestris is Latin and means “Alps”, “from the Alps”. This species is characterised by a bluish dorsal surface and a vivid orange ventral surface that runs down to the groin (Griffiths, 1996; Gasc et al., 1997). They have smooth, slimy skin in the aquatic phase, while their skin becomes granular, dry, rough and impervious in the terrestrial phase. The alpine newt has a small head. Their tails are thin, flattened and usually equal to the length of their body, including the head. They have robust limbs, silvery eyes and two lines of vomerine teeth. Alpine newts lack webbed hind feet (Griffiths, 1996; Arnold, 2003). Sexual differentiation can be done using morphometric traits; in particular, they are very easy to tell apart during the breeding season (aquatic phase) (Arnold, 2003). The males have a white/silver longitudinal band running along the lower part of the flank with distinctive black speckles, while the females lack this band. During the aquatic phase, males possess a low light blue tail crest that is covered with dark blue/black spots as well as a short, white body crest with the same design as on the lower flanks. Females lack these crests. In general, the males have a bluish appearance with light shades, while the females have brown skin and can even be black in colour (Arnold, 2003; Wells, 2007). Since the dorsal crests are involved in the males’ complex courtship behaviour, they become particularly noticeable during the breeding season (Griffiths, 1996). Unlike salamanders, newts generally spend the majority of their life in the water; however, most European species live on the land and only spend the breeding season in the aquatic phase (Gasc et al., 1997). Ichthyosaura alpestris inhabits from sea level up to 2,500 m. In the aquatic phase, they prefer stagnant waters such as shallow ponds, temporary pools, lakes, ditches, slow-moving streams and drinking troughs. In the terrestrial phase, the alpine newt is nocturnal by nature. However, they are diurnal in the breeding season (Kuzmin,

15 INTRODUCTION

1995; Gasc et al., 1997). Hibernation can occur both on land and in the water. Depending on the altitude, breeding season occurs from the middle of February/March (Spellerberg, 2002). Although no risks pose a major threat to the alpine newt in Belgium, they are currently listed as a vulnerable species in Brussels (Kestemont, 2010).

1.2.2.2. Lissotriton helveticus, Palmate newt Lissotriton helveticus (Figure 2B), formerly Triturus helveticus, is the smallest native newt in Belgium. Lissotriton is originates from Greek and is composed of two words: Liss means “smooth” and Triton means “newt”. Helveticus originates from Latin and means “Swiss”. The base colour of both sexes is light brown or olive green. Tail length is slightly shorter than snout-vent length. Glandular ridges alongside the back give the male newts a square-backed appearance. A dark, bandit-like line runs horizontally from the nostrils, across the head and along the eyes. They have a yellow/orange belly, which can sometimes be spotted (Griffiths, 1996; Gasc et al., 1997). The tail has an orange line along the centre. The males and some of the females have a dark, spotted pattern on the dorsolateral surfaces. In the terrestrial phase, the skin traits change and become almost black in colour with dry, rough and impervious integument (Nöllert and Nöllert, 1992; Gasc et al., 1997). Sexual differentiation is easily discerned through morphological traits, particularly in the aquatic phase. During breeding season, the hind limbs of males become quite webbed and look like a palm leaf. Moreover, the male’s tail crest becomes tall and dark brown with a fine filament at the end, while females have a low tail crest without a filament. In the terrestrial phase, the two ridges running along the males’ backs give them a box-like appearance (Arnold, 2003; Staniszewski, 2011). Although the palmate newt is superficially similar to the smooth newt and might be mistaken for them, the throat of the palmate newt is plain pink or yellow and never spotted. This key characteristic is used to distinguish between the two species (Griffiths, 1996). The altitudinal distribution ranges from sea level up to 1500m (Gasc et al., 1997). The aquatic phase occurs in shallow ponds with calm waters. This species has a distinct preference for oligotrophic conditions and slightly acidic water with a low accumulation of dissolved salts (Nöllert and Nöllert, 1992; Wells, 2007). Their terrestrial habitat is composed of upland areas and lowland heaths such as moorland, prairies and forests (Gasc et al., 1997). Depending on the latitude and altitude, hibernation starts in November and lasts until late February/March. Palmate newts usually hibernate on land, in sheltered spots like beneath stones. During the breeding season, they migrate to the aquatic site as early as February and

16 INTRODUCTION mating takes place in aquatic conditions (Griffiths, 1996). In the breeding season, they become active in both the day and night. However, they are active during rainy and humid nights during the terrestrial phase (Griffiths, 1996; Gasc et al., 1997). Lissotriton helveticus is currently under substantial threat in Belgium and is listed as a rare species in Flanders, a near-threatened species in Wallonia and a vulnerable species in Brussels (Bauwens and Claus, 1996; Kestemont, 2010).

1.2.2.3. Lissotriton vulgaris, Smooth newt Lissotriton vulgaris (Figure 2C), formerly Triturus vulgaris, is the most widespread newt species of the Lissotriton genus in Europe (Griffiths, 1996). Lissotriton: refers to Lissotriton helveticus. Vulgaris originates from the Latin for “common” (Gasc et al., 1997). The dorsolateral colouration can be pale brownish, greyish-brown or yellowish-brown covered with rounded, dark green or brown speckles. The ventral surface is yellow/orange with dark spots. The throat is conspicuously spotted. This trait is used for distinguishing the smooth newt from the palmate newt, whose throat is not spotted (Gasc et al., 1997; Arnold, 2003). In contrast to the female, the male’s cloaca is swollen and noticeable. Moreover, the males have a single black stripe along the centre of the spine, while the females have paired parallel stripes on each side of the spine (Gasc et al., 1997; Staniszewski, 2011). It is easy to distinguish between males and females during the aquatic phase. A continuous, high serrated body and tail crest covered with dark speckles emerges on the dorsal surface and concurrently, a light bluish marking appears on both side of the tail in males, while the female’s crest is not developed. Males possess developed webbing on their hind limbs. In addition, the males are brownish in colour with many round speckles, while the females are a uniform, brown colour or sometimes greenish (Griffiths, 1996; Gasc et al., 1997). Smooth newts can be found up to an altitude of 1,000 m (Nöllert and Nöllert, 1992). They are primarily terrestrial, except during the breeding season, and select a variety of habitats spread across varied regions such as bush lands, deciduous woodlands, marshes and anthropogenic landscapes like parks, fields and gardens. In addition, they can come into the steppe zone by dwelling in wooded river valleys (Gasc et al., 1997; Wells, 2007). The Smooth Newt spends most of the time on land. It returns to water bodies for reproduction in spring or autumn, depending on the geography and the subspecies (Griffiths, 1996). Adult newts are active primarilyin darkness, both in water and on land. In the daytime, terrestrial adults are active during and just after rains. In the aquatic phase, adults and larvae are active both day and early night. During metamorphosis, larval activity shifts to the

17 INTRODUCTION twilight (Nöllert and Nöllert, 1992). Although the smooth newt is listed as safe/low risk species in Belgium, their populations are currently declining. This species is listed as a vulnerable species in Wallonia and Brussels (Kestemont, 2010).

1.2.2.4. Salamandra salamandra, Fire salamander Salamandra salamandra (Salamandra is the Greek word meaning “salamander”) (Figure 2D) is easily identified by the dorsolateral black colour covered with large and vivid yellow to red blotches and/or spots along the back and flanks. These features can vary among subspecies, even within the same population. (Nöllert and Nöllert, 1992; Gasc et al., 1997). Salamandra salamandra subspecies terrestris is found in Belgium. This subspecies is characterized by the yellow dorsal spots, which are often arranged to form two lateral lines. The stripes can be continuum or interrupted (Griffiths, 1996). The parotoid glands are noticeable and located behind the pigmented eyes. The ventral surface is black or dark brown with or without markings. The length of the tail is shorter than the body, including the head. They lack crests as well as webbing on the limbs (Staniszewski, 2011). Females are generally larger than males and possess relatively shorter extremities and tail. The male's cloaca is much more swollen than the female's cloaca (Gasc et al., 1997). The fire salamander inhabits regions below 2,200 m in altitude that are associated with wet and cool forests, often in beech forests and limestone areas close to streams and rivers. The occurrence of certain populations on woodless hillsides likely indicates the previous existence of forests. Due to a lack of webbed limbs and poor swimming skills, the fire salamander is not found in floodable regions (Nöllert and Nöllert, 1992; Gasc et al., 1997). Salamanders are nocturnal. However, during rainy weather, they regularly leave their hiding places by day. Mating occurs on land and mostly in spring or autumn (Griffiths, 1996). Since this species is viviparous, females move to the water after mating and release the larvae into well-oxygenated water and fishless parts of brooks (Gasc et al., 1997, Arnold, 2003; Wells, 2007). The number of larva varies between 8 and 55 for Salamandra salamandra subspecies terrestris. Larval development takes several months, but in many cases they overwinter and finish their metamorphosis in the next year (Gasc et al., 1997). Recently, a steep decrease in fire salamander populations that has brought this species to the edge of local extinction has been observed in the Netherlands (Martel et al., 2013). Due to a reduction in the population density of fire salamanders, this species is currently considered to be vulnerable in Belgium (Bauwens and Claus, 1996; Kuijken, 1999).

18 INTRODUCTION

1.2.2.5. Triturus cristatus, Great crested newt Triturus cristatus (Figure 2E) is a warty newt. Triturus originates from Greek and is composed of two words: the first (Triton) comes from Greek mythology about the sea guardian and the second (ura) means “tail”. Cristatus originates from Latin and means “crest”. The great crested newt is characterised by greyish/brownish to black, rough skin covered with small, whitish, wart-like points on the flanks (Griffiths, 1996). The ventral surface is yellow/orange with black speckles. The males have a very large and serrated body and tail crests, the colour of which is the same dark colour as the dorsolateral surface. These two crests are distinctly separated (Gasc et al., 1997). Like all true newts, this species lacks parotoid glands. The tail is long and approximately equal to the body length, including the head. It has symmetrical vomerine teeth (Nöllert and Nöllert, 1992). There are significant sexual differences in seasonal variation. In the aquatic phase, males possess a deeply notched middorsal crest and a non-jagged crest on the tail, while females do not have crests. Moreover, males have a blue-white flashing along the lateral surface of the tail, while females have a reddish/orange band along the underside of the tail. During the terrestrial phase, the males’ crests become indistinctive and the skin turns out to be black in colour in many specimens, while females have a light orange stripe along the middorsal surface (Gasc et al., 1997; Staniszewski, 2011). Terrestrial habitation occurs up to 1,100 m in altitude including in woodlands, scrub and rough grasslands near the aquatic breeding sites (Griffiths, 1996). The aquatic habitats are made of deep, vegetated pools, large water areas and slow moving rivers, and sometimes in eutrophic water bodies. Triturus cristatus is more dependent on suitable ponds than most other newts of the genus Triturus and Lissotriton in Western Europe (Gasc et al., 1997; Wells, 2007). Depending on the latitude and altitude, hibernation starts in October/November and can last until February/March. Hibernation sites consist of damp places such as deciduous woodland floors amid fallen branches and leaves (Griffiths, 1996). On returning to the aquatic sites, mating is started once the breeding characteristics of males are completed. The great crested newt is nocturnal by nature in both its terrestrial and aquatic habitats (Nöllert and Nöllert, 1992). In Flanders, this species is still present in all provinces of Flanders, but scattered over very fragmented population. The same holds true for Wallonia where the species is declining but still present in all major ecoregions (Louette and Bauwens, 2013).

19 INTRODUCTION

Figure 2. Native Caudata : Ichthyosaura alpestris (A), Lissotriton helveticus (B), Lissotriton vulgaris (C), Salamandra salamandra (D), Triturus cristatus (E). Photos credited by Frank Pasmans.

2. Invasive amphibians 2.1. Invasive species: definition An invasive amphibian species also referred to as an alien, non-native or non- indigenous species is a species that has been introduced outside its natural geographic range. The introduction of invasive alien species can occur in a natural environment accidently or deliberately, mainly due to human interference because of the increasing trade in amphibians around the world (Lowe et al., 2000). The invasion process in a new environment by an alien species occurs in the following steps (Groves, 1986; Richardson et al., 2000): I) Introduction: As a result of human activity, either deliberate or accidental, non-native species arrive to a new geographical region. A non-native species can be successfully introduced after overcoming the geographical barriers and survive in a new environment beyond their former geographical zone. The new species within this phase is called an exotic species. II) Acclimatisation: When environmental barriers do not prevent new amphibian species from surviving, they form a colony in the new environment. The new species within this step is termed an acclimatised species. III) Naturalisation: This step occurs when new species overcome reproductive barriers and have normal reproduction in the wild. Thus, they have ability to breed and produce fertile offspring over several lifecycles without direct intervention by humans. The new species within this step is called a naturalised species.

20 INTRODUCTION

IV) Expansion: When naturalized species overcome dispersal barriers, they sustain new self- perpetuating populations with widespread dispersal and undergo incorporation within the new habitat. The new species within this phase is termed invasive species (Groves, 1986; Richardson et al., 2000). Invasive species can cause enormous ecological damages if released or transported to a different environment (Rosen and Schwalbe, 1995). The presence of bioinvaders can lead to the restructuring of established food webs, the predation on native species, the introduction of new diseases to the new surroundings, and competition with indigenous animals for space and food. Other ecological changes occur when the non-native species interbreed with native species, likely altering the gene pool. This may lead to hybridization, which reduces biodiversity and the ecosystem's ability to adapt to natural or human-induced changes (Drake and ICSU; 1989; Cox, 1999; Hanselmann et al., 2004; Garner et al., 2006). Since the invasion stage of alien species within a specific country determines the priority actions to prevent introduction and modify the adverse impacts, invasive species are classified in the following list (ISEIA, 2009): I) Alert list species: The alert list consists of the invasive species that are rare to non- existent within a specific country, but which are invasive in neighbouring regions and can be recognised as a potential threat to native amphibians. II) Species under naturalisation (isolated populations): Species that have reached the prime stage of the invasion process within a specific country. They only establish small, isolated populations located in the vicinity of their introduction points. III) Naturalised species with restricted range (restricted populations): populations of species that overcome barriers to successful breeding and start the active distribution phase, but of which the dispersion is still restricted to some specific region within a specific country. IV) Widespread naturalised species: populations of species that have already overcome all biogeographic barriers to establish widely distributed, self-perpetuating populations within a specific country.

2.2. Invasive amphibians of Belgium There are two invasive species recognised as being at different invasion stages in Belgium: Lithobates catesbeianus and Pelophylax ridibundus (Table 3; Figure 3). In addition to these widespread invasive species, two other exotic species i.e. Italian crested newt (Triturus carnifex) and African clawed toad (Xenopus laevis) are also introduced to northern Europe. These introductions may lead to colonies in Belgium. (Alien Invaders-Europe

21 INTRODUCTION

Introduced Exotics, 2013).

Table 3. Invasive amphibian species and invasion stage in Belgium Scientific name Common name Family Range Since Lithobates catesbeianus American bullfrog Ranidae Restricted 1996 Pelophylax ridibundus Marsh frog Ranidae Restricted 1975

Figure 3. Invasive species: Lithobates catesbeianus (A), Pelophylax ridibundus (B). Photos credited by Frank Pasmans.

2.2.1. Lithobates catesbeianus, American bullfrog Lithobates catesbeianus (Figure 3A) is native to the north-eastern United States and inhabits areas from the east coast to the Rocky Mountains (Govindarajulu et al., 2006). Lithobates originates from Greek and is composed of two words: “litho” means “stone” and “bates” denotes “to walk”. Catesbeianus is dedicated to Catesby, who discovered this species in the Carolinas, in the United States. They are considered to be one of the most detrimental alien invasive species and hold a spot on the list of “100 of the world’s worst invasive alien species” (Lowe et al., 2000). They play a significant role in the deterioration of amphibian populations, a phenomenon that has been observed all over the world (Daszak et al., 2004; Hanselmann et al., 2004; Garner et al., 2006). Their presence can negatively alter the biodiversity of native amphibian species, mostly through competition for food resources and predation of other amphibian species (Kiesecker and Blaustein, 1997, 1998; Kats and Ferrer, 2003). They can also transmit pathogens such as Batrachochytrium dendrobatidis (causative agent of chytridiomycosis) and ranaviruses to native amphibians (Hanselmann et al., 2004; Garner et al., 2006). The American bullfrog larvae are highly competitive with native amphibian larvae and can inhibit the growth and development of native species by decreasing the food supply (Jooris, 2005). In addition, their skin excretions can curtail growth

22 INTRODUCTION and even reduce the survival rate percentage in other amphibian species (Laufer and Sandte, 2004). American bullfrogs can be easily distinguished from the native frogs of Belgium through their large body size. In addition, the presence of conspicuous supratympanic folds behind the eyes and absence of dorsolateral folds makes them easy to distinguish from the European Pelophylax (Jooris, 2002 a, b). The average male snout-vent length is 152 mm and 162 mm for females (reaching up to 200 mm long) and grow up to 800 g in weight. Their tadpoles are also very large and measure up to 140 mm (Jooris, 2002 a, b). American bullfrogs have a wide, flat head with a rounded snout, very large tympanums and horizontal pupils. Their skin is quite smooth. The hind limbs possess well-developed webbing. They lack dorsolateral folds and parotoid glands (Jooris, 2002 a, b). American bullfrogs have an internal vocal sac opening at the corner of the mouth. The dorsal colouration varies between light olive green to quite dark with dark brown blotches in a net-like pattern along the back. The ventral surface is whitish, grey or yellowish, with or without darker spots. They usually have green heads with golden eyes (Mudde, 1992). Males are smaller than females. Mature males have large tympanums that are twice the diameter of the eye, while mature females have eardrums that are about the same diameter of the eye. In the breeding season, males possess a yellow throat, while the females have a dirty white throat mottled with dark speckles. Males have darkly pigmented nuptial pads (Stebbins, 2003). They are active by day and night (Jooris, 2005). They hibernate from October and usually bury themselves in the mud or in leaf litter. The breeding season begins in spring and lasts throughout early summer, but can vary according to latitude (Durham and Bennett, 1963). The new tadpoles emerge between July and September and they usually complete the stages of metamorphosis after two years, but sometimes they can live up to three or four years before turning into frogs (Jooris, 2005). Tadpoles are mainly herbivorous, although they sometimes feed on invertebrate prey; adult American bullfrogs are generalist predators that have a wide diet including invertebrates, amphibians, fish, small rodents, snakes, reptiles and birds (Beringer and Johnson, 1995). Juvenile American bullfrogs can migrate up to a maximum recorded distance of 1,600 meters when seeking shelter in the vegetation and food; however, adult American bullfrogs usually stay close to the pond or along the shoreline between hibernation and the breeding season (Ingram and Raney, 1943). American bullfrogs can occupy any aquatic habitats that are below an altitude of 50 meters, such as lakes, streams, marshes, ponds (irrigation and brackish), ditches and backwaters (Durham and Bennett, 1963). Most amphibian species are found in temporary

23 INTRODUCTION ponds, but the American bullfrog typically inhabits permanent ponds (Jooris, 2005). Adult American bullfrogs are often found in warmer wetlands with intermediate minimum annual temperatures and high precipitation, such as still waters with vegetation, and they are not present in areas with very cold winters (-20˚C) (Ficetola et al., 2006). They also tend to prefer artificial and human-modified habitats like fish ponds. Human-driven habitat modification such as changes in hydrology from seasonal to permanent water, shifting of emergent vegetative cover and increasing water temperature can create suitable habitats for American bullfrogs (Hayes and Jennings, 1986; Doubledee et al., 2003). American bullfrogs were mainly imported to Europe for breeding in frog farms to harvest them for human consumption and for garden ponds in the pet trade (Daszak et al., 2006). Since the 1930s, the American bullfrog has been the subject of at least 25 independent introductions in Europe in 8 countries, including Belgium, France, Germany, Greece, Italy, the Netherlands, Spain, and the United Kingdom (Lanza, 1962; Stumpel, 1992; Ficetola et al., 2006). Although the introduction of the American bullfrog has been forbidden in European countries (law of the European Council 2551/1997) since 1997, approximately 60% of the introductions had already occurred during the 1980s and 1990s (Jooris, 2005; Ficetola et al., 2006). The first and largest acclimatisations were recorded in northern Italy, with smaller, isolated populations along the western coast of Italy (Albertini and Lanza, 1987). In 1990, they established a population through successful reproduction in northwestern Europe (Stumpel, 1992). The American bullfrogs were probably introduced to Belgium in two main ways, i.e. primary introduction from North America on a large scale for the pet trade and secondary translocations (accidentally or deliberately) by humans from neighbouring countries (Stumpel, 1992; Ficetola et al., 2006, 2007). Subsequently, the successful acclimatisation of American bullfrogs in Belgium has been made possible due to the presence of several nutrient-rich, lentic habitats (e.g. lakes and large, deep ponds) within close proximity to each other (Stumpel, 1992; Ficetola et al., 2006). Successfully established populations can be found in both Flanders and Wallonia (Jooris, 2002a; Ficetola et al., 2006).

2.2.2. Pelophylax ridibundus, Marsh frog Pelophylax ridibundus (Figure 3B) has been successfully established in Belgium since the 1970s (Hulselmans, 1978; Burny and Parent, 1985). Pelophylax: refer to Pelophylax lessonaes. Ridibundus is from Latin and means “to laugh in bursts” or “to burst with laughter”, illustrative of their call. Marsh frogs are classified into a genus of green frogs that includes to the edible frog (Pelophylax kl. esculentus) and the pool frog (Pelophylax

24 INTRODUCTION

lessonae). Although Pelophylax ridibundus might have invaded Belgium by natural range expansion from neighbouring countries, Holsbeek et al., (2008) suggested that its invasion is at least partly enhanced by commercial trade, with origins as far as the Middle East. Although distinguishing between the three species of water frog may be difficult, the average snout-vent length of the marsh frog (up to 140 mm) is larger than that of the other two native water frogs in Belgium (Hulselmans, 1978). Males have a pair of noticeable vocal sacs, which are grey in colour. The marsh frog has a proportionally longer heel on the hind leg than the two other native Pelophylax and can extend this up to the eye (Baloutch and Kami, 1995; Tarkhnishvili and Gokhelashvili, 1999). The dorsal pattern varies widely between specimens and can be uniform grey in colour to yellowish/green, with or without dark blotches. There is often a light middorsal stripe running along the back. The ventral surface is whitish and mottled with darker spots (Baloutch and Kami, 1995). Marsh frogs possess a moderately sharp snout, prominent dorsolateral folds, well-developed webbing on the hind feet, vomerine teeth, and golden eyes with horizontal pupils. The posterior part of the tongue is free and forked (Baloutch and Kami, 1995; Tarkhnishvili and Gokhelashvili, 1999). The eyes are located on top of the head, which allows the marsh frog to keep just its eyes above the water’s surface, while their body is safely submerged. They lack parotoid glands (Kuzmin, 1999). Differentiation between the sexes is possible because the males have a pair of external vocal sacs behind the corners of the mouth and dark nuptial pads on the first finger. The male’s nuptial pads turn darker in the breeding season in particular. In addition, males are often smaller than females and may be two thirds the size of the females (Baloutch and Kami, 1995; Tarkhnishvili and Gokhelashvili, 1999). The marsh frog is a highly opportunistic species and can be found in all types of bodies of water, including flowing and stagnant water habitats such as dams, lagoons, lakes, rivers and waterways. They prefer quite warm or temperate areas with rich, herbaceous vegetation (Hulselmans,1978; Burnyand and Parent, 1985). Marsh frogs show a strong ecological competition with other native water frogs, particularly with the indigenous edible frogs. In addition, they are described as a potential threat at the genetic level for populations of native species (Holsbeek and Jooris, 2009). Depending on the elevation, hibernation occurs from September/October to the beginning of June (Kuzmin, 1995). As a rule, hibernation occurs in water, but in some locations, it occurs in rodent burrows and holes in river banks and lake shores (Gasc et al., 1997). Group hibernation is typical, but the groups usually do not exceed several dozen individuals. Breeding starts from several days to one month after the frogs' spring appearance

25 INTRODUCTION

(Baloutch and Kami, 1995). This species has succeeded in establishing a self-perpetuating population in both Flanders and Wallonia over the last two decades (Jooris, 2002; Percsy, 2002).

3. Amphibian Diseases Several studies have reported population declines, range reduction and extinction of amphibian species around the world (Collins et al., 1988; Cunningham et al., 1996; Greer et al., 2005; Schock and Bollinger 2005, Teacher, 2008; Ariel et al., 2009). Amphibians are currently considered to be the most threatened vertebrate class on the planet (Gascon et al., 2007). There are numerous reasons cited for the dramatic decline in amphibian populations: I) Changes to habitats, including deforestation, wetland removal, habitat fragmentation and global warming, have significantly affected amphibian populations (Vos and Chardon, 1998; Kolozsvary and Swihart, 1999; Carey and Alexander, 2003; Collins and Strofer, 2003; Delis et al., 2004; Silvano and Segalla, 2005; Raffel et al., 2006). II) Human activities, such as harvesting amphibians from wild populations, can significantly deplete the target species from a given region (Stuart et al., 2004). The removal of amphibians from wild populations for many purposes, such as food for human consumption, for alleged medicinal and scientific purposes, and for commercial reasons, e.g. the pet trade, has a negative impact on amphibian populations (Costa-Neto, 2004; Schlaepfer et al., 2005; Humraskar and Velho, 2007; Nickerson and Briggler, 2007). It appears that Ranidae (true frogs) have been overwhelmingly affected by human activities, whereas Bufonidae, Leptodactylidae and Hylidae (tree frogs) are being devastated by habitat loss (Stuart et al., 2004). III) The introduction of an exotic species to a habitat can adversely affect indigenous amphibians (Moyle, 1973; Bradford, 1989; Fisher and Shaffer, 1996; Crossland, 1998). Typically, Salmonid fish, bullfrogs and pantropical introduction of marine toads (Bufo marinus) have been detrimental to native amphibians because these prey upon native amphibian larvae and compete for food resources (Moyle, 1973; Hammerson, 1982; Jennings and Hayes, 1985; Hayes and Jennings, 1986; Crossland, 1998, 2000; Adams, 1999). IV) The Earth’s thinning ozone layer may lead to increased ultraviolet radiation of amphibians and their larvae resulting in severe damage and population declines (Middleton et al., 2001; Blaustein et al., 2003; Hatch and Blaustein, 2003). Although increased ultraviolet light has been demonstrated in some regions experiencing

26 INTRODUCTION

amphibian decline, the degree to which amphibians are affected has not yet been ocumented (Corn and Muths, 2002; Palen et al., 2002). V) Acidification of both terrestrial waters and soils has negatively affected all types of amphibians resulting in a weakening of the natural immune system (Vatnick et al., 2006). Acidification is typically carried by small airborne particles and wet deposition

containing acidic industrial by-products such as sulphur dioxide (SO2), nitrogen oxide

(NOx) and ammonium (NH4). Amphibian larvae are particularly sensitive to a change in the pH of their aquatic habitat (Dunson et al., 1992; Preest, 1993). VI) Contaminating the environment with a wide range of chemical substances (e.g. nitrate fertiliser and heavy metals) has also adversely affected amphibian populations (Hatch and Blaustein, 2003; de Wijer et al., 2004; Chen et al., 2006, 2007). VII) Recent research has identified two diseases that have been implicated in global increase in amphibian mortality: chytridiomycosis, an emerging fungal disease caused by the pathogens B. dendrobatidis and Batrachochytrium salamandrivorans, and ranaviral disease caused by ranaviruses (Collins and Storfer 2003, Daszak et al., 2003, Martel et al., 2013). These diseases are now globally notifiable to the World Organisation for Animal Health (OIE). Environmental stressors such as metal disorders (deficiency/overloading) may also be a concern, particularly in combination with diseases; they can be lethal to already weakened individuals (Stuart et al., 2004; Wake and Vredenburg, 2008). Therefore, understanding and controlling diseases has become a major focus of both in situ and ex situ amphibian conservation efforts worldwide (Gascon et al., 2007). This part of the dissertation reviews the biology of the causative agents of ranavirosis and chytridiomycosis, the clinical signs of these diseases, the routes of transmission and the currently known pathogenesis.

3.1. Ranavirosis 3.1.1. Background The family Iridoviridae is classified into five genera, i.e. Chloriridovirus, Iridovirus, Lymphocystivirus, Megalocytivirus and Ranavirus, and several tentative species such as Singapor grouper iridovirus (SGIV) and Grouper iridovirus (GIV) (Williams, 1996; Chinchar et al., 2009; ICTV, 2012). They can infect both invertebrates (Chloriridovirus and Iridovirus) and ectothermic vertebrates (Lymphocystivirus, Megalocytivirus and Ranavirus) and have a global distribution (Chinchar et al., 2009). Four recognised species in the genus Ranavirus

27 INTRODUCTION can infect amphibians, including frog virus 3 (FV3), Ambystoma tigrinum virus (ATV), Bohle iridovirus (BIV), and Rana catesbeiana virus Z (Chinchar et al., 2009). Ranaviruses have been detected worldwide and have caused significant morbidity and mortality in amphibian species over the past two decades (Williams et al., 2005). They can infect both wild and captive amphibians from at least 14 families and over 70 individual species on 5 continents (except Antarctica and Africa), with most reported cases occurring in the family Ranidae (Miller et al., 2011). The first reported FV3 isolation from amphibians occurred in 1966 in leopard frogs (Lithobates pipiens, formerly Rana pipiens) being examined for Lucke’s renal adenocarcinoma (Granoff, 1966). In Europe, ranaviruses were first isolated by Fijan et al. (1991) during an investigation into Pelophylax esculentus mortality in Croatia. Prior to this, a viral haemorrhagic septicaemia had been reported in Pelophylax esculentus (formerly Rana esculenta) in the former Yugoslavia, while ranaviruses was assumed to be the causative agent (Kunst and Valpotic, 1968). In 1993, the first unambiguous mortalities due to ranavirus infections reported in common frogs (Rana temporaria) in the southeast of England (Cunningham et al., 1993). Subsequently, many outbreaks have occurred in both wild and captive amphibians across Europe (Table 4). Since the 1980s, ranaviruses isolates have markedly increased, which has resulted in ranaviruses being described as one of the emerging pathogens in amphibians (Daszak et al., 1999). Recently, the World Organisation for Animal Health placed Ranavirus infections as notifiable diseases for amphibians (OIE, 2012).

28 INTRODUCTION

Table 4. Summary of countries and amphibian families in which Ranavirus infection has been reported in Europe Country Family Scientific Name Virus W, C* References

Tylototriton Pasmans et al., Belgium Salamandridae FV3 kweichowensis C 2008 Croatia Ranidae Pelophylax esculentus Unclassified Ranavirus W Fijan et al., 1991 Denmark Ranidae Pelophylax esculentus Unclassified Ranavirus W Ariel et al., 2009 the Ranidae Pelophylax spp. W CMVT Kik et al., 2011 Netherlands Salamandridae Lissotriton vulgaris W Alves de Matos et Portugal Salamandridae Triturus marmoratus Unclassified Ranavirus W al., 2008 Alytidae Alytes obstetricans W Balseiro et al., Spain CMTV Salamandridae Ichthyosaura alpestris W 2010 Pelophylax esculentus C Switzerland Ranidae FV3 Miller et al., 2011 Pelophylax ridibundus C Alytidae Alytes obstetricans W Duffus et al., 2009 Hyatt et al., 2000, Salamandridae Lissotriton vulgaris W 2010 the UK Unclassified Ranavirus Cunningham et Bufonidae Bufo bufo W al., 1993, 1996 Ranidae Rana temporaria W Drury et al., 1995 * W=wild population, C = captivity including zoological and ranaculture facilities.

3.1.2. General characteristics Morphologically, ranaviruses are large with a distinctive icosahedral symmetry (125-300 nm in diameter; Williams et al., 2005). The virion possesses an internal lipid membrane between the viral core and outer capsid, while the capsid itself is covered by an external membrane with a series of proteins. They have linear, double-stranded DNA genomes (105 kbp) with a GC content of 55% (He et al., 2002; Tan et al., 2004). These genomes are circularly permuted and terminally redundant (Goorha and Murti 1982; Delius et al., 1984). They are found in an enveloped or non-enveloped (naked) form, although both forms are infectious (Chinchar et al., 2009). However, enveloped virions are more infectious, which may be due to at least two different host receptor proteins (Williams et al., 2005). Enveloped virions enter cells via receptor-mediated endocytosis and are uncoated in lysosomes, while naked virions uncoat at the plasma membrane (Braunwald et al., 1985). Typically, the FV3 genome (105 kbp) encodes 98 non-overlapping, open reading frames (ORF). These genes encode transcription and express factors leading to the

29 INTRODUCTION appearance of immediate early (IE), delayed early (DE), and late (L) viral transcripts (Majji et al., 2009). IE and DE genes are responsible for nucleic acid metabolism and immune evasion, while L genes are responsible for DNA packaging and virion assembly (Chinchar et al., 2009; Majji et al., 2009). Although the cellular receptor for the entrance of FV3 is highly conserved among vertebrate hosts, encompassing mammals, birds, reptiles, and fish (Chinchar et al., 2009), temperatures above 32°C do not permit viral replication. Thus, the in vivo host ranges are restricted to poikilothermic vertebrates (Chinchar et al., 2009).

3.1.3. Replication cycle Naked virions release the DNA core into the host cell cytoplasm by binding to the cell membrane, while enveloped virions enter the host cell through receptor-mediated endocytosis (Williams et al., 2005). The main events in ranavirus replication are illustrated in Figure 4.

Figure 4. Ranavirus replication cycle (adapted from Williams et al., 2005)

The DNA core is translocated from the cytoplasm to the nucleus, where IE transcription and viral mRNA synthesis using host RNA polymerase II (Pol II) takes place (Chinchar et al., 2009). Following synthesis of the viral IE transcripts and translation, IE protein(s) set the

30 INTRODUCTION stage for the transcription of DE genes (Chinchar et al., 2009), which encodes catalytic proteins, such as the viral DNA polymerase (Lua et al., 2005). Subsequently, the viral materials, including IE and DE transcripts in the nucleus, commence the first stage of DNA synthesis using viral DNA polymerase (Chinchar et al., 2009). Newly synthesised viral DNA is transported to the cytoplasm where it serves in the formation of concatameric (multiple repeats) structures (Goorha, 1982). This viral DNA is methylated by a virus-encoded DNA methyltransferase (Chinchar et al., 2009). DNA methylation is crucial to viral proliferation because it blocks the induction of pro-inflammatory responses by blocking toll-like receptor 9 (TLR); treatment with a DNA methylation (5-azacytidine) inhibitor reduces virus production 100 times (Chinchar et al., 2009). Despite this, the mechanisms of virion assembly are poorly known (Chinchar et al., 2009); at least twelve viral genes are involved in virion assembly (Chinchar and Granoff, 1984). Following late viral RNA transcription and protein synthesis, virions are assembled in distinct areas of the cytoplasm called “viral assembly sites” (AS) (Chinchar et al., 2009). Some of the main structural elements, such as the major capsid proteins, are encoded by late viral proteins (Zhao et al., 2007; Chinchar et al., 2009). The virions are packaged by a headful mechanism resulting in a terminally redundant and circularly permuted virion (Goohra and Mutri, 1982; Chinchar et al., 2009). Subsequently, naked virions accumulate in paracrystalline arrays within the cytoplasm (Chinchar et al., 2009). Some virions acquire an envelope as they bud from the plasma membrane; however, the majority of virions remain cell-associated upon lysis of the infected cell (Chinchar et al., 2009).

3.1.4. Transmission and pathogenesis Transmission can predominantly occur among amphibian populations through multiple horizontal routes (direct or indirect), including exposure to contaminated water or fomites, ingestion of infected tissue -either through necrophagy or cannibalism, and direct contact with infected individuals (Wolf et al., 1969; Jancovich et al., 1997; Pearman et al., 2004; Brunner et al., 2004, 2005, 2007). Vertical transmission may occur through the exposure of the eggs to an infected cloaca during oviposition or through infected sperm during fertilisation (Gray et al., 2009). Mechanical transmission by insects and leaches has also been suggested, but it needs more investigation (Converse et al., 2005; Miller et al., 2007). Ranaviruses have inhibitory effects on the synthesis of cellular macromolecules (protein, RNA and DNA) following productive infection (Chinchar et al., 2009).

31

INTRODUCTION

Interestingly, treatment of infected cells with heat-inactivated or UV-inactivated viruses indicates that viral structure proteins play a major part, rather than a newly synthesised viral gene product (Chinchar et al., 2009). In addition to the inhibition of cellular synthetic functions, the two mechanisms - including necrosis and apoptosis - are assumed to lead to cell death within 6-9 hours post- infection (Chinchar et al., 2009). However, apoptosis through DNA fragmentation, chromatin condensation and membrane reversal is currently considered to be the predominant mechanism (Chinchar, 2003; Chinchar et al., 2009). The pathogenicity of ranaviruses is affected by different factors in amphibians, including species susceptibility, host immune responses, and environmental stressors (Hoverman et al., 2010). Susceptibility to ranaviruses differs within amphibian species as well as across phylogenetic lineages (Schock et al., 2010; Hoverman et al., 2011). The variation of susceptibility is associated with intrinsic factors, suggesting that the loss of genetic diversity and adaption of host immune evasion genes to local ranaviruses strains coincides with increasing mortality risk (Pearman et al., 2005; Ridenhour et al., 2008). However, phylogeny is still an important factor affecting a species’ susceptibility to ranavirus infection. Species within the family Ranidae are more susceptible to infection compared to species in the families Ambystomatidae and Hylidae (Hoverman et al., 2011). The susceptibility to ranaviruses also varies between amphibian developmental stages, even in the same species (Gray et al., 2009). Two critical periods of infectivity have been recognised at developmental stages, i.e. the time before the larval immune system develops, and during metamorphosis while the larval immune system is being reorganised. Therefore, the amphibian’s susceptibility to ranaviruses might be increased at those times. Despite several reports indicating that recent metamorphs, are the most susceptible age groups in North America, adults are more commonly reported in die-offs of several amphibians species in Europe (Ariel et al., 2009; Duffus et al., 2010; Hoverman et al., 2010; Robert, 2010; Teacher, 2010; Miller et al., 2011). This difference may be a consequence of population density, resulting in increasing transmission and elevating viral titers in adults, while undeveloped immune responses may be associated with high mortality in recently metamorphosed juveniles (Robert et al., 2005; Green et al., 2010). Although the embryos are susceptible to ranaviruses, the egg appears to protect the embryos from infection due to the mucopolysaccharide/mucoprotein capsule coating its surface or the vitelline membrane around the embryo (Tweedell et al., 1986; Duffus et al., 2008).

32 INTRODUCTION

Surprisingly, virulence is considerably correlated with the viral strain and not the amphibian lineage (Brunner and Collins, 2009). In addition, the effect of temperature on pathogenesis has been demonstrated through both virus-specific factors or/and host factors, e.g. robust immune responses at higher temperatures (Rojas et al., 2005). Temperatures at or near 18°C appear to be optimum for ATV to infect and rapidly kill Ambystoma tigrinum. The mortality of salamanders at 18°C appears to be a result of the interaction of rapid viral multiplication with reduced host immune response at this temperature. At colder temperatures (10°C), time to death is greater but virus titer per animal is increased (Rojas et al., 2005). Innate immunity plays a distinct protective role in susceptible species through the secretion of anti-microbial peptides (AMP) on amphibian skin. These components can significantly reduce viral infectivity by incapacitating the virus (Chinchar et al., 2001, 2004; Rollins-Smith, 2009). Peritoneal leukocytes, natural killer cells and T-cells can also establish a critical, innate defence mechanism against ranaviruses in amphibians (Morales et al., 2010). Potent adaptive immunity might also protect them by inducing a CD8 T-cell response and IgY that can protect the amphibian upon re-infection with ranaviruses (Gantress et al., 2003; Robert, 2010). Environmental stressors such as a high density of animals, anthropogenic stressors and toxicants lead to a higher mortality in amphibians because the energetic resources are diverted away from the immune response in order to cope with the stressors (Forson et al., 2006; Echaubard et al., 2010; Hayes et al., 2010). The geographic elevation of populations may also factor into host susceptibility, indicating that morbidity of infection increases at higher elevations (Gray et al., 2007; Gahl et al., 2008).

3.1.5. Clinical signs and pathology Ranavirus infections can cause a variety of symptoms in the various growth stages of amphibians. Affected larvae display behavioural changes, such as abnormal swimming patterns, buoyancy problems, lordosis and abnormal posture, while adults often exhibit lethargy, anorexia or are simply found dead (Bollinger et al., 1999; Densmore and Green, 2007). Based on the developmental stages of being infected by ranaviruses, two forms of ranavirosis can be described in Anurans: the acute and the chronic form (Duffus and Cunningham, 2010). Acute ranavirosis, or systemic haemorrhage syndrome (HS), can often demonstrate in larvae (Duffus and Cunningham, 2010). During this phase, several types of haemorrhages, such as ecchymosis near the vent, petechiation and/or ecchymosis of internal organs - especially the mesonephros of kidneys and liver, can affect the tissue (Cunningham

33

INTRODUCTION et al., 1996, 2008). In addition to HS, edema and swelling can be observed in multiple areas of the body (Cunningham et al., 2008; Gray et al., 2009). Moreover, irregular patches can occur due to loss of pigmentation in the skin (Duffus et al., 2010). The chronic form, ulcerative syndrome (US), is characterised by cutaneous erosions and ulcerations of the skin of the hind limbs and ventral pelvic region and may persist over days or weeks (Cunningham et al., 1996, 2007, 2008). US primarily appears in metamorphs and adult Anurans in Europe (Duffus et al., 2010). In some cases, necrosis can appear in the digit(s) or extremities of the limbs, resulting in loss of the digit(s), especially in regions with harsh winters (Cunningham et al., 1996). The gall bladder may also be enlarged, but this is attributed to anorexia (Miller et al., 2011). Adult Caudata demonstrated haemorrhages occurring on the plantar surface of the feet and tail and edema appearing in the gular fold (Docherty et al., 2003; Converse et al., 2005). Polypoid lesions can occur in the early stages of the disease in Eastern tiger salamanders (Ambystoma tigrinum) and can progressively cover most of the body (Jancovich et al., 1997). Histological findings between adult and juvenile amphibians are similar and reveal multiple necrotic foci scattering within tissues. However, the rates of necrotic foci in internal tissues of animals with the cutaneous form are less extensive (Cunningham et al., 1996). Virus particles (intracytoplasmic) comprising primarily basophilic inclusion bodies have been inconsistently detected in different cells/tissues, including epithelial cells, renal tubular cells, hepatocytes, meninges, the neuroepithelium, nasal tissues, adipose tissue, the trachea, muscles, and osteoclasts (Jancovich et al., 1997; Docherty et al., 2003; Gray et al., 2009; Miller et al., 2011). However, virus particles can infrequently appear as eosinophilic inclusions (intranuclear), which are attributed to the stage (early versus late) of infection (Miller et al., 2011). Virus particles can cause organ failure within a few days to a few weeks of exposure (Brunner et al., 2005; Forson et al., 2006). Since the intestinal epithelial cells are the primary route of entry into the body, these cells appear to be one of the first cell types to be infected, as early as 3 hours post-exposure (Miller et al., 2007; Robert et al., 2011). Subsequently, the kidney (renal tubular epithelium) is considered to be the target organ, especially in immunocompromised amphibians (Robert, 2010). Microscopic changes can be challenging due to primary viral lesions compounded by opportunistic bacteria (Cunningham et al., 1996). Several recorded observations have provided evidence of subclinical infections in larvae. These findings indicate that asymptomatic carriers of ranaviruses can occur often in amphibian populations (Brunner et al., 2004; Harp et al., 2006; Gray et al., 2007).

34 INTRODUCTION

3.1.6. Diagnosis Several diagnostic methods have been identified, including histopathology, viral isolation, ELISA, microscopy, molecular techniques and cytology (Hyatt et al., 2000; Gray et al., 2009). A conventional PCR test is performed as both a screening and diagnostic test to identify ranavirus infections in blood and tissue (fresh, frozen, formalin-fixed, or paraffin-embedded tissues) (Green et al., 2009). A 500 base pair region of the major capsid protein (MCP) is amplified during the test. This region is a highly conserved segment of the ranaviruses genome without any other homologous sequences in the NCBI GenBank database (Mao et al., 1996; Greer and Collins, 2007). This test is highly specific for ranaviruses, but the sensitivity of the test depends on the type of sample collected and the time since the amphibians were exposed to the virus (Greer and Collins, 2007). The tail clip versus whole-body homogenate has less probability of contacting a virus particle, and amphibians may also yield a low level of virus in the early stages of infection, resulting in a false negative result, which may the researcher to underestimate the true prevalence of the virus (Greer and Collins, 2007). Since a positive PCR result only indicates the presence of virions and does not confirm an active infection, supportive tests such as a histological examination must be performed to differentiate between ranavirus infection versus a disease. A quantitative PCR (qPCR) test based on the viral DNA polymerase gene can detect lower amounts of genetic content than the conventional PCR, and consequently, it may be more sensitive than the conventional PCR (Gray et al., 2009; Holopainen et al., 2011). The immunohistochemistry (IHC) methods can detect the viral antigens in formalin- fixed tissues through anti-ranavirus antibodies. Although IHC methods can definitively demonstrate systemic ranavirus infection in different amphibian tissues, these are only used in the detection of, and not in the differentiation between ranaviruses (Holopainen et al., 2009; Cinkova et al., 2010). Preferably, IHC methods should be used at the outbreak of ranaviruses (Gray et al., 2009). Serologic assays can detect antibodies against ranaviruses in amphibians; in particular, ELISA can detect BIV antibodies in cane toads (Bufo marinus) (Whittington and Speare, 1996; Whittington et al., 1997). Due to the occurrence of several cross-reactions in polyclonal antibody-based methods, these methods can also be used in the detection of, but not the differentiation between ranaviruses (Ahne et al., 1998; Hyatt et al., 2000; Bang Jensen et al., 2009; Gobbo et al., 2010). Virus isolation in cell culture is considered to be one of the most important methods for the detection of ranaviruses, however, it cannot demonstrate active infection (Gray et al., 2009; Holopainen et al., 2009). Several

35 INTRODUCTION fish cell lines, such as fathead minnow epithelium and papilloma cyprinid cells, are currently available. Since virus isolation is dependent on different factors, such as tissue type, degree of post-mortem autolysis and the ability to optimise the incubation conditions for a specific strain, infection cannot be excluded from consideration based only on negative isolation results (Cullen and Owens 2002). Electron microscopy (EM) can be used for visualisation of the icosahedral-shaped virions within formalin-fixed or/and paraffin-embedded tissues as well as for confirmation of a cultured virus (Gray et al., 2007; Burton et al., 2008; Miller et al., 2009). While the previous diagnostic methods are used to detect ranaviruses, a histological examination is specific in confirming the presence of disease by demonstrating haemorrhage, cellular necrosis and intracellular inclusion bodies (Miller et al., 2009). Subsequently, IHC and/or EM can support the histological finding (Gray et al., 2007).

3.1.7. Conservation strategies Although there is no single cause for amphibian decline, ranavirus infections result in at least localised die-offs of amphibians (Gray et al., 2007). Conservation strategies must therefore be implemented to reduce ranaviruses disease emergence (Gray et al., 2007).

I) Strategies that influence pathogen abundance and transmission: The possibility of exposure to ranaviruses is largely affected by the environmental prevalence of virions (Gray et al., 2007; Blaustein et al., 2012). Since the virus is inactivated following pond drying (Brunner et al., 2007), amphibian species that breed in ephemeral ponds versus permanent bodies of water may be exposed to ranaviruses less frequently (Blaustein et al., 2012). Thus, pond drying can be considered to be a possible form of prevention of ranaviruses exposure within natural ponds. However, inter- and intraspecific reservoirs and frequency-dependent transmission in an amphibian may maintain the virus in the habitat between years (Gray et al., 2007). Therefore, regional transport of live amphibians must be accompanied with certification of ranaviruses-negative test results (Gray et al., 2007). Field findings reveal that equipment from field workers such as clothes and vehicles can potentially spread ranaviruses between areas (Langdon, 1989). Therefore, hygiene methods must improve in order to reduce this potential risk, e.g. by applying a 3% bleach or 0.75% Nolvasan (2% chlorhexidine diacetate; Fort Dodge Animal Health) solution for 1 minute (Bryan et al., 2009).

36 INTRODUCTION

II) Strategies that influence host responses to infection: Anthropogenic stressors can negatively impact amphibian responses to ranaviruses infection and contribute to disease dynamics within their habitats (Blaustein et al., 2012). Both fertilisers and pesticides can potentially increase susceptibility to ranaviruses by compromising amphibian immunity (Davidson et al., 2002; Forson and Storfer, 2006). Additionally, cattle farming is known to negatively influence amphibian populations due to the increase of nitrogenous compounds in the water resulting in immunosuppression in the amphibian (Jancovich et al., 1997; Gray et al., 2007; Greer and Collins, 2008). Therefore, strategies to minimise anthropogenic stressors must be implemented by establishing suitable vegetation buffers around breeding sites and restricting the use of chemicals in amphibian breeding sites (Semlitsch and Bodie 2003; Gray et al., 2007).

3.2. Chytridiomycosis 3.2.1. Background Chytridiomycosis is caused by two chytrid fungi: B. dendrobatidis and B. salamandrivorans (Berger et al., 1998; Martel et al., 2013). This fungal disease is described as the worst infectious amphibian disease, driving them to extinction (Gascon et al., 2007). Currently, B. dendrobatidis infects over 350 amphibian species in 37 countries on all continents except Antarctica and is considered to be a pandemic pathogen (Fisher et al., 2009; Kriger and Hero, 2009). It has been also implicated as the driving force behind the decline of more than 200 amphibian species (Skerratt et al., 2007). In some cases, chytridiomycosis is described as being responsible for the extinction of entire species in the wild such as gastric brooding frogs (Rheobatrachus sp.), and the sharp-snouted day frog (Taudactylus acutirostris) (Retallick et al., 2004; Schoegel et al., 2005; Gewin, 2008). B. salamandrivorans causes erosive skin disease and rapid mortality in fire salamanders (Salamandra salamandra), both in the wild as under experimental conditions.This fungus was only very recently discovered and was implicated in a dramatic population decline among fire salamanders in the Netherlands (Martel et al, 2013). The actual distribution of this fungus, its pathogenesis and possible treatment remain to be studied. Together with the closely related B. dendrobatidis, this taxon forms a well-supported chytridiomycete clade, adapted to vertebrate hosts and highly pathogenic to amphibians. However, the lower thermal growth preference of B. salamandrivorans, compared with B. dendrobatidis, and resistance of midwife toads (Alytes obstetricans) to experimental infection with B. salamandrivorans suggest differential

37 INTRODUCTION niche occupation of the two chytrid fungi (Martel et al., 2013). In 1998, B. dendrobatidis was first identified in wild frogs in Queensland (Australia) and Panama during mass mortality events (Berger et al., 1998). In 1999, it was detected in a captive blue poison dart frog (Dendrobates azureus) at the Smithsonian National Zoological Park, Washington, D.C., United States (Longcore et al., 1999). Subsequently, B. dendrobatidis has been associated with devastated amphibian populations, severe population declines affecting multiple species extinctions occurring around the world, notably in Central America, Australia, as well as in Europe (Berger et al., 1998; Bosch et al., 2001; Lips et al., 2006; Skerratt et al., 2007; Bielby et al., 2009; Walker et al., 2010). In 2008, B. dendrobatidis was listed as a notifiable pathogen by the World Organization for Animal Health (OIE, 2008). B. dendrobatidis is widespread across Europe. Eleven out of 15 countries have infected amphibians in the wild. However, previous estimates of prevalence surely underestimated how heavily B. dendrobatidis had penetrated Europe. Prevalence at population levels in Switzerland, Austria, France and the UK were all seriously underestimated in the earlier study [respectively, Switzerland current estimate, 54% (unpublished), previous estimate 25%; Austria current 20% (Sztatecsny and Glaser, 2011), previously 0%; France current 34% (unpublished), previously 0%; UK current 26% (unpublished) previously <1%]. Other national estimates currently available include Spain (~25%; Walker et al., 2008) and the Netherlands/ Belgium (~30%). The overall prevalence for Europe stands at 27% (1351 positives/ 4999 samples) (Fisher et al., 2012). In Belgium, clinical chytridiomycosis was first detected in three captive Central American plethodontid salamanders (Bolitoglossa dofleini) (Pasmans et al., 2004). Subsequently, a high number of B. dendrobatidis organisms was reported in a dead indigenous common midwife toad (Alytes obstetricans) (Pasmans et al., 2010). The percentage of B. dendrobatidis-positive samples in different European countries is shown in Figure 5.

38 INTRODUCTION

Figure 5. B. dendrobatidis distribution in wild populations across Europe (adapted from RACE, 2013)

The origin of B. dendrobatidis has yet to be identified. However, epidemiological evidence provides that B. dendrobatidis could have originated in Africa, Asia (Japan) or eastern North America, and was disseminated across different geographic regions through anthropogenic movement of amphibians for laboratory research subjects, as pets, as educational or display animals and as part of conservation and breeding programmes (Weldon et al., 2004; Garner et al., 2006; Goka et al., 2009; Soto-Azat et al., 2010). This has led to international World Organisation for Animal Health Recommendations, which require that amphibians have to be free of any B. dendrobatidis infection before international shipment (Schloegel et al., 2010). Due to the global emergence of chytridiomycosis since the 1990s, two hypotheses were proposed to explain B. dendrobatidis epidemiology, i.e. the novel or spreading-pathogen hypothesis (NPH) and endemic pathogen hypothesis (EPH) (Skerratt et al., 2007; Fisher et al., 2009). The NPH states that the spread of B. dendrobatidis is a result of its introduction into naïve host populations through the amphibian trade (Skerratt et al., 2007). On the other hand, the EPH states that B. dendrobatidis was present in global amphibian populations, but population declines were caused by changes in host susceptibility, pathogen virulence, environmental changes, or a combination of these factors (Morgan et al., 2007; James et al., 2009). Although genetic similarity among geographically disparate isolates provides evidence to support the NPH, recently developed markers showed few alleles per loci and revealed that it has yet to delineate spatial population structure (Morehouse et al., 2003; Morgan et al.,

39

INTRODUCTION

2007; James et al., 2009). In addition, the supposition that it spreads as a wave front supports the novel pathogen hypothesis in Australia, Central and South America (Laurance et al., 1996; Berger et al., 1998; Skerratt et al., 2007; Lips et al., 2006, 2008). Because of the emergence of B. dendrobatidis, even in wilderness regions, the chytrid-thermal-optimum hypothesis (CTOH) has also been proposed (Bosch et al., 2007). In this hypothesis increasing temperatures are proposed to have increased B. dendrobatidis growth that then led to the observed population declines (Pounds et al., 2006; Bosch et al., 2007). However, recent investigations suggested that temperature can be considered to be a correlative factor rather than a causative one (Rohr et al., 2008). A combination of both the NPH and EPH is currently believed to have caused the pandemic chytridiomycosis in amphibians (Goka et al., 2009; Walker et al., 2010). Therefore B. dendrobatidis can be introduced into new habitats by a NPH-like process and be expressed consistently by an EPH-like environmental context.

3.2.2. Phylogenetic characteristics of B. dendrobatidis Based on the zoospore ultrastructure as well as DNA analysis of the small subunit ribosomal DNA (SSU-rDNA), B. dendrobatidis is phylogenetically placed in the order Rhizophydiales (phylum Chytridiomycota, class Chytridiomycetes) (James et al., 2006; Letcher et al., 2006). The term “Chytrid” fungi derived from a Greek word, “chytridion”, meaning pot or container, and refers to approximately 1,000 heterotrophic species of several phyla, whereas only two species, namely B. dendrobatidis and B. salamandrivorans, are recognised as the causes of the disease (chytridiomycosis) in amphibians (Berger et al., 1998; Longcore et al., 1999; Pasmans et al., 2006, Martel et al., 2013).

3.2.3. Lifecycle stages B. dendrobatidis has two known life stages that are typical for chytrids: a sessile, reproductive thallus with a single zoosporangium and motile, uniflagellated zoospores released from the zoosporangium (Berger et al., 2005). Zoospores develop into germlings by encysting and resorbing the flagellum and then a monocentric thallus structure appears. The thallus grows larger and becomes multinucleate through mitotic divisions, resulting in zoosporangium over 4 to 5 days at 22°C (Berger et al., 2005). Germination of a zoospore cyst or germling is followed by the development of a germ tube that invades the epidermal cell of tolerant or susceptible species. At the end of the germ tube, a swelling is formed that gives rise to a new thallus. Then, cell contents of the germling

40 INTRODUCTION migrate into the newly formed thallus and the emptied germling evanesces. The new intracellular thallus forms a rhizoid-like structure that extends to a deeper epidermal layer and develops a swelling at its end and a new intracellular thallus is formed (Greenspan, 2012; Van Rooij et al., 2012) (Figure 6).

Figure 6. Schematic representation of the B. dendrobatidis lifecycle in susceptible host (A) and tolerant host (B). The endobiotic lifecycle in (A) includes successively germ tube mediated invasion, establishment of intracellular thalli, spread to the deeper skin layers, upward migration by the differentiating epidermal cell to finally release zoospores at the skin surface. The epibiotic lifecycle in (B) includes germ tube mediated invasion, outgrowth of a rhizoidal network, uptake of host cell cytoplasma as nutrient for the growing and maturing chytrid thallus upon the skin surface. Adapted from Van Rooij et al. (2013).

A colonial thallus can also be the result of the formation of more than 1 zoosporangium from 1 zoospore and is considered to be the only known variation of the cycle (Longcore et al., 1999). The released zoospores can swim up to 24 hours, covering a distance of 2 cm in still media (Berger et al., 2005). Optimal growth occurs between 17-25°C and at a pH of 6-7. However, B. dendrobatidis can grow and reproduce in vitro under a range of temperatures (4- 25°C) and pH levels (4-8) (Berger et al., 2005).

3.2.4. Susceptibility to B. dendrobatidis Amphibian species differ with regard to their susceptibility to B. dendrobatidis (Woodhams et al., 2006b; 2007a; Searle et al., 2011b). Host response to infection can be roughly categorized as resistant (fast clearance of the infectious agent), tolerant (persistent infection in absence of disease) and susceptible (infection resulting in lethal disease)

41

INTRODUCTION

(Schneider and Ayres, 2008; Raberg et al., 2009). For instance, two species, i.e. the Western toad (Bufo boreas) and the mountain yellow- legged frog (Rana muscosa), show a 100% mortality rate due to B. dendrobatidis infection, while the Eastern tiger salamander (Ambystoma tigrinum) appears to be resistant to B. dendrobatidis infections (Davidson et al., 2003; Carey et al., 2006; Rachowicz and Briggs, 2007). Some other amphibians such as the American bullfrog (L. catesbeianus) and the African clawed frog (Xenopus laevis) can act as carriers of B. dendrobatidis due to their low level of infection (Skerratt et al., 2007). Moreover, within a single species some populations coexist with B. dendrobatidis without no evidence of disease while others go extinct (Pilliod et al., 2010; Puschendorf et al., 2011). Different developmental growth stages of amphibian also differently respond to the pathogen. Larvae are likely to be much more resistant compared with juveniles and adults, which may suffer high mortality (Briggs et al., 2010; Kilpatrick et al., 2010). Since immune responses undergo a dramatic reorganisation at metamorphosis and the post-metamorphic mechanisms are not yet complete, newly metamorphic animals are considered to be individuals that are vulnerable to disease (reviewed by Rollins-Smith, 1998). Therefore, B. dendrobatidis infections can frequently cause substantial mortality in the post-metamorphic phase of many species (Bosch et al., 2001; Briggs et al., 2005; Carey et al., 2006; Garner et al., 2009). B. dendrobatidis strains are significantly different in their virulence and at least three divergent lineages, i.e. Global Panzootic lineage (BdGPL), Swiss lineage (BdCH) and Cape lineage (BdCAPE) show variable virulence (Farrer et al., 2011). The global panzootic lineage has emerged across at least five continents during the 20th century and is associated with the onset of epidemics in North America, Central America, the Caribbean, Australia, and Europe (Farrer et al., 2011). Interestingly, virulence of the pathogen can itself be influenced by environmental factors, resulting in varying avirulent to virulent states (Bosch et al., 2007; Fisher et al., 2009). Environmental factors contributing to susceptibility of amphibians to this disease are not well known. Temperature and geographic elevation have been suggested as key factors since chytridiomycosis is thought to occur more frequently at cool high-elevation sites where enigmatic amphibian declines have been observed (Young et al., 2001; Lips et al., 2006; Smith et al., 2007; Fisher et al., 2009). In addition, infection prevalence and severity exhibit strong seasonality, increasing in the cooler months of early spring (Bosch et al., 2001; Drew et al., 2006; Kriger et al., 2007). Moreover, changes in biodiversity can have several outcomes on disease risk, including

42 INTRODUCTION dilution and amplification effects, both of which can have a profound influence on the effects of disease in a community (Searle et al., 2011a). Searle and co-workers experimentally manipulated host diversity and density in the presence of B. dendrobatidis and found a dilution effect where increased species richness reduced disease risk, even when accounting for changes in density (Searle et al., 2011a). These results demonstrate the general importance of incorporating community structure into studies of disease dynamics and have implications for the effects of B. dendrobatidis in ecosystems that differ in biodiversity (Searle et al., 2011a).

3.2.5. Clinical signs, gross pathology and pathogenesis Clinical signs of chytridiomycosis may include abnormal posture, such as splayed limbs, tetanic spasms upon handling, depression, loss of righting reflex, excessive shedding of the skin, reddening of the skin and lethargy in juveniles and adult frogs (Simoncelli et al., 2005; Cunningham et al., 2005; Berger et al., 2009). However, most of the clinical signs are non-specific for this disease (Cunningham et al., 2005; Berger et al., 2009). In tadpoles, only the mouthparts are keratinized and susceptible to B. dendrobatidis infection (Berger et al. 1998), leading to mouthpart depigmentation and sometimes defects (Marantelli et al., 2004). B. dendrobatidis can infect both the superficial (stratum granulosum and stratum corneum) and deeper epidermal cells of amphibians. They initially grow in living cells, but the spherical sporangia occur within the superficial cells of the keratinised epidermis and release zoospores into the environment through the skin’s surface (Berger et al., 2005). Therefore, chytridiomycosis represents a sustained cutaneous infection in amphibians (Berger et al., 1998). Histologically, B. dendrobatidis can cause two key lesions in the epidermis based on an amphibian’s developmental stages: first, mouthpart abnormalities resulting in the loss of sections of the keratinised mouthpart in tadpoles, leading to emaciation; and the second, hyperplasia in the stratum granulosum and parakeratotic hyperkeratosis in the stratum corneum of skin cells particularly on the belly, digits, and pelvic patch resulting in the sloughing off of surface layers in post-metamorphic animals (Berger et al., 1998, 2005; Fellers et al., 2001). Moreover, haemorrhages in the skin, muscles, or eyes; hyperaemia of the skin on the digits and ventrum, especially the hyper-vascularised pelvic patch known as the “drink patch”; congestion of viscera; epidermal sloughing; and rare epidermal ulcers might be observed in infected animals (Berger et al., 1998; Daszak et al., 1999; Simoncelli et al., 2005).

43

INTRODUCTION

Although the pathogenesis of B. dendrobatidis is not yet well understood, recent studies indicate that electrolyte depletion plays a critical role in the pathophysiology of chytridiomycosis (Voyles et al., 2012). The normal osmoregulation and ion balance across the skin is disrupted through the breakdown of transport processes in the infected amphibian’s skin (Voyles et al., 2009; Carver et al., 2010). In severe cases of chytridiomycosis, death by asystolic cardiac arrest can occur due to reductions in the electrolyte concentrations of the blood plasma (hyponatraemia and hypokalaemia) (Voyles et al., 2009, 2012).

3.2.6. Transmission Since B. dendrobatidis zoospores have a distinctive flagellum, they can spread by swimming through water or moist environments (Berger et al., 1998). The amphibian skin subsequently comes into contact with water or moist substrates, such as wet soil or equipment containing zoospores. B. dendrobatidis can also infect amphibians through direct contact between an infected animal and an uninfected one during territorial or breeding encounters (Berger et al., 2009; Johnson and Speare, 2005). Water birds and insects have been identified as possible vectors (Cashins, 2009; Garmyn et al., 2012). Also crayfish contribute to the dissemination of B. dendrobatidis, as this fungal pathogen can survive on crayfish, causing gill pathology and mortality (McMahon et al., 2013).

3.2.7. Amphibian defense against chytridiomycosis Defense mechanisms against chytridiomycosis have been extensively reviewed by van Rooij (2013). Relatively little is known with regard to innate and acquired immune responses against chytridiomycosis. First line defense mechanisms possibly include the amphibian cutaneous mucus layer mainly composed of mucin glycoproteins, antimicrobial peptides (AMPs) and antifungal metabolites produced by the skin microbiome (reviewed in Rollins- Smith and Conlon, 2005; Rollins-Smith et al., 2009). Mucus is secreted from mucous glands and mainly consists of glycosylated mucins and mucopolysaccharides. The mucin composition not only determines the viscosity of mucus, but also its capacity to harbor and retain components secreted by amphibian skin such as antimicrobial peptides (AMPs) and antibodies (Rollins-Smith and Conlon, 2005; Richmond et al., 2009; Ramsey et al., 2010). Skin mucus by itself forms a mechanical barrier, hindering pathogens access to cell surfaces (Ramsey et al., 2010). AMPs are secreted from granular glands secrete upon irritation, stress or adrenergic stimulation. About forty AMPs have been shown to inhibit B. dendrobatidis in vitro (Rollins-

44 INTRODUCTION

Smith et al., 2009). Subsequently, the suite of antimicrobial peptides expressed by each species may predict whether they survive or decline when confronted with natural B. dendrobatidis infections in the wild. Species expressing a mixture of skin peptides that potently inhibits B. dendrobatidis growth in vitro are among the more common and resistant species. In contrast, declining or endangered species express peptides with poor in vitro anti- B. dendrobatidis activity (Woodhams et al., 2006b). The metabolites 2,4-diacetylphloroglucinol (2,4-DAPG), indol-3-carboxaldehyde (I3C) and violacein secreted by symbiotic skin bacteria, such as Janthinobacterium lividum (Harris et al., 2006; Brucker et al., 2008) have been shown to inhibit B. dendrobatidis in vitro and in vivo (Harris et al., 2006; Woodhams et al., 2007a; Brucker et al., 2008, Lam et al., 2010; Flechas et al., 2012). For instance, pretreatment of Rana muscosa juveniles with Janthinobacterium lividum protected them from lethal B. dendrobatidis infections that occurred in untreated but B. dendrobatidis-exposed frogs (Harris et al., 2009). Also components of the adaptive immune system e.g. immunoglobulins (Ig) IgM, IgX and IgY (Ramsey et al., 2010) found in skin secretions are thought to contribute to resistance/ tolerance by binding to and eliminating B. dendrobatidis. Although the data support a role for the adaptive immune system in resistance of some species to B. dendrobatidis, immunization with heat-killed B. dendrobatidis to induce a protective systemic immune response did not protect boreal toads (Bufo boreas) or mountain yellow-legged frogs (Rana muscosa) when they were exposed to relatively high numbers of infectious zoospores (Rollins-Smith et al. 2009; Stice and Briggs 2010). This suggests that conventional immunization protocols that induce elevated serum antibodies would not necessarily be protective against this pathogen, which is confined to the skin.

3.2.8. Diagnosis B. dendrobatidis infections can be demonstrated through different methods such as histopathology, immunohistochemistry (IHC), and PCR (Berger et al., 1998, 2002; Briggs and Burgin, 2003; Van Ells et al., 2003; Olsen et al., 2004; Boyle et al., 2004). Histopathology can be used in the early stages of infection, when the zoosporangia are formed within the cells of the stratum granulosum and stratum corneum. For this purpose, samples are taken from toe clips and ventral skin and fixed in 10% neutral buffered formalin or 70% alcohol. Examination of gross sections with hematoxylin and eosin stain can be done (Hyatt et al., 2007; Berger et al., 2009). Although spherical or oval zoosporangia (5 to 13 μm) with a

45

INTRODUCTION thin eosinophilic wall can be seen by light microscope, the presence of zoosporangia can be confirmed using special fungal stains such as a periodic acid-Schiff (PAS) or silver stain (Hyatt et al., 2007; Berger et al., 2009). The different contents of the zoosporangia, including immature sporangia with central basophilic homogenous masses, or multinucleate and empty sporangia with the presence of discharge tubes, can be explained by the lifecycle of the fungus (Berger et al., 2005, 2009). Focal hyperkeratosis and hyperplasia are described as common histological findings (Berger et al., 1998, 2009). Since the organism can be easily missed, especially in light infections, this method has no high sensitivity (Van Ells et al., 2003). Immunohistochemical staining (using polyclonal antibodies against B. dendrobatidis in an immunoperoxidase test) is a useful screening method for B. dendrobatidis when combined with morphological findings and symptom identification (Berger et al., 2002). It can detect the pathogen with slightly more increased sensitivity than the histopathological stain and is more specific than special fungal stains (Berger et al., 2002; Hyatt et al., 2007; Skerratt et al., 2011). Two molecular assays, i.e. a conventional PCR and a quantitative real-time PCR, can be broadly used to detect the pathogen (Annis et al., 2004; Boyle et al., 2004). The real-time q-PCR is the most common diagnostic test and carried out by taking a non-invasive swab sample from the tadpole’s mouthpart or the adult’s ventral surface. Due to high sensitivity of q-PCR, this test can thus detect the intensity of infection and demonstrate the presence of carriers (Hyatt et al., 2007). Confirmation of chytridiomycosis is reported by using a combination of clinical signs, histopathological findings and pathogen detection by PCR. Although the pathogenic burden can vary between species, approximately 10,000 zoospore equivalents on PCR is an indication of severe to lethal disease in amphibians (Kinney et al., 2011).

3.2.9. Preventative management strategies Although there are no proven methods to control the disease in the wild, general hygiene and biosecurity practices aim to mitigate the spread of chytridiomycosis into naïve amphibian populations (Phillott et al., 2009). The development of preventative strategies can be compiled using the following methods:

I) Developing trade and quarantine regulations: According to the NPH, B. dendrobatidis has been widely introduced into new habitats through the anthropogenic movements of amphibians (Skerratt et al., 2007). Thus, quarantine and movement limitations should be implemented to protect uninfected habitats from Bd-

46 INTRODUCTION infected regions. Amphibians should be placed in a different container when shipping between countries. All newly acquired, captive amphibians should be placed in quarantine initially for at least 2 months, keeping the temperature between 17 and 23°C to increase the chance of infection becoming clinically apparent (Lynch, 2001; Daszak et al., 2001). During the quarantine period, individuals should be regularly examined for skin swabs for PCR assay (Young et al., 2007). All water, soil, plants, litter or any other wet objects that came into contact with the amphibian that was imported into a country, or moved between locations within a country, should be disinfected using effective chemical substances (i.e. didecyldimethylammonium chloride at >0.0012% final concentration for 2 minutes, or sodium hypochlorite at >1% for 1 minute) or thermal sterilisation by immersion in hot water (>47°C for 30 minutes or 60°C for ≥5 minutes), which will kill B. dendrobatidis (Johnson and Speare, 2003; Young et al., 2007). Other effective hygiene methods include using disposable gloves and changing them between enclosures, using automated husbandry systems, disinfecting the water used in the enclosure before disposal, disinfecting footwear and equipment between the sites, and attending to animals in a consistent order, starting with threatened species and those least likely to be infected (Marantelli and Berger, 2000; Lynch, 2001).

II) Developing strategies for captive management and conservation: When B. dendrobatidis infection is prevalent in captive collections, elimination of the pathogen is crucial, not only for the health of captive species, but also to remove the pathogen from the environment in which amphibians may be reintroduced (Martel et al., 2011). Itraconazole has been used successfully both orally (2-13 mg/kg daily for 9–28 days) and via shallow immersion (0.01% suspension for 5 minutes daily for 11 days) to treat some adult amphibians (Taylor et al., 1999; Nichols & Lamirande, 2000; Taylor, 2001). Voriconazole (sprayed once daily for 7 days with a 1.25 μg/ml solution in water) is also considered to be one of the most effective treatments for captive amphibians (Martel et al., 2011). Although thermal treatment, with the increase of the temperature up to 37°C, can be an effective treatment, most species do not tolerate this high temperature (Woodhams et al., 2003; Young et al., 2007). Amphibian skin harbours symbiotic resident microbes, which constitute the only line of defence that is not directly host produced and has been successfully manipulated to mitigate disease (Harris et al., 2009; Vredenburg et al., 2011). Growing evidence supports the hypothesis that antifungal skin microbes suppress chytridiomycosis (Harris et al., 2009;

47 INTRODUCTION

Vredenburg et al., 2011; Muletz et al., 2012). Bacteria inhibit B. dendrobatidis growth directly through production of inhibitory metabolites and perhaps indirectly through immunomodulation where microbes regulate the production of host defenses such as antimicrobial peptides (AMPs) and lysozyme (Reid et al., 2011). Importantly, a field experiment involving bioaugmentation of an anti-B. dendrobatidis species, Janthinobacterium lividum, on Rana mucosa in the Sierra Nevadas showed that frogs treated with a probiotic bath had lower peak infection loads than untreated controls. One year after treatment, untreated control were not recovered, suggesting that probiotic treatment allowed individuals to persist by preventing B. dendrobatidis from reaching a lethal threshold (Vredenburg et al., 2011). Therefore, areas still naïve to B. dendrobatidis provide an opportunity for conservationists to proactively implement bioaugmentation strategies to prevent further amphibian declines. In areas where B. dendrobatidis is endemic, bioaugmentation can facilitate repatriation of susceptible amphibians currently maintained in assurance colonies (Harris et al., 2009; Schmeller et al., 2014).

3.3. Zoonotic bacterial diseases 3.3.1. Chlamydia The causative agents belong to the order of the Chlamydiales and are coccoid, obligate intracellular pathogens (Bodetti and Timms, 2002). Chlamydiosis is a serious infection of amphibians and the disease also affects humans, birds, and mammals (Schacter et al., 1999). Based on histologic lesions and the presence of inclusion bodies, these infections were originally attributed to Chlamydophila psittaci in amphibians (Mutschmann, 1998). C. psittaci can be transmitted to humans and many other mammals and can cause Psittacosis as severe enteric and respiratory illness in many avian species and humans (Schacter et al., 1999). Using molecular methods such as PCR, it has since been shown that other species of Chlamydiales have been associated with these infections, including C. pneumoniae, C. abortus, and C. suis (Berger et al., 1999; Blumer et al., 2007). Moreover, a novel species of Chlamydiales named Candidatus Amphibiichlamydia salamandrae has recently been found in diseased salamanders (Martel et al., 2012). The pathogenicity of this novel species in humans has not been demonstrated yet and probably it has a restricted host range. Chlamydiosis was originally recognized in a mass mortality of African clawed frogs (Xenopus laevis) fed uncooked beef livers (Reed et al., 2000). Infected frogs may die peracutely or exhibit lethargy, dysequilibrium, cutaneous depigmentation, petechiae, and edema (Berger et al., 1999). Histologically, intracytoplasmic basophilic inclusion bodies can

48 INTRODUCTION be identified in sinusoidal lining cells of the liver and spleen (Newcomer et al., 1982; Howerth, 1984). Secondary bacterial infections are frequently present in affected amphibians and must be treated appropriately. Antibiotic treatment including doxycycline (5–10 mg/kg/day, PO) or oxytetracycline (50 mg/kg/day, PO) may be effective against chlamydial infection (Pessier, 2002).

3.3.2. Salmonella Salmonella are members of the large bacterial family Enterobactericae (Le Minor, 1991). The presence of Salmonella is well documented in amphibians, particularly in anurans (Ang et al., 1973; Chambers and Hulse, 2006; Drake et al., 2013). In caudata, Salmonella infections probably only exceptionally occur (Pasmans et al., 2013). None of 100 faecal samples from captive urodelans from three different collections yielded any Salmonella isolates (F. Pasmans, unpublished results). Possibly, the low body temperatures of most urodelan species (<20°C and even <15°C) may limit colonization by Salmonella. This finding is in support of the hypothesis that ectothermic vertebrates with low body temperatures are not suitable hosts for Salmonella bacteria. Although pathogenic Salmonella species may be carried by amphibians, they rarely cause to appear clinical signs of disease and it seems to be of little significance to the amphibian host (Reichenbach-Klinke and Elkan 1965; Anver and Pond 1984). Prevalence of Salmonella isolated in surveys from clinically normal amphibians is generally greater than 10% and sometimes as high as 60%, particularly if intestinal contents are sampled at several sites (Sharma et al., 1974). The isolation of bacteria in high concentrations in intestinal contents of toads suggested that anuran amphibians might be good reservoirs of Salmonella with a high potential to contaminate the environment and humans (Sharma et al., 1974; O'Shea et al., 1990). Salmonella are excreted in faeces by amphibians, and can cause severe disease in humans and domestic animals (Hemingway et al., 2009). For humans salmonellosis is a notifiable disease due to its potential severity, but also because it is a marker of the microbiological safety of the human food chain. Water borne infections can occur, but are unusual (Taylor et al 2000). An outbreak of gastroenteritis in humans was thought to have possibly arisen from green tree frogs (Litoria caerulea) contaminating drinking water in rainwater tanks (Taylor et al., 2000). Salmonellosis associated with amphibian contact has been also described previously (e.g. Srikantiah et al., 2004), one notorious outbreak, caused by S. Typhimurium, involving at least 85 people from 31 US states (CDC, 2010). This

49 INTRODUCTION outbreak was linked to contact with water of aquaria containing African dwarf clawed frogs (Hymenochirus sp.). The consumption of contaminated frog legs also poses a public health risk (Purushothaman et al., 1981; Catsaras, 1983; Rajagopalan et al., 1983).

3.4. Mesomycetozoans Mesomycetozoans are members of an unusual clade of eukaryotic protists that exist phylogenetically near the animal-fungal divergence (Densmore and Green, 2007). Multiple mesomycetozoan genera, including Amphibiothecum (formerly Dermosporidium), Amphibiocystidium, and Ichthyophonus, are known pathogens of amphibians. Nomenclatural changes based on molecular characterization of the mesomycetozoans is ongoing, therefore many of these organisms are in the process of being taxonomically reorganized (Feldman et al., 2005). Recent information supports the designation of the genera Amphibiothecum and Amphibiocystidium to include amphibian pathogens formerly in the genera Dermocystidium, Dermosporidium, and Dermomycoides (Pascolini et al., 2003). Various species of Amphibiocystidium and Amphibiothecum are known to affect a variety of anurans including bufonids, ranids, and hylids and also salamanders. The organisms are spore-forming pathogens that occur in cysts, which are typically located in the ventral dermis. The infections are generally self-limiting and nonlethal, healing within 4 to 8 weeks after the development of clinical signs. Grossly, infections appear as small to large (> 1 cm) multifocal nodules or pustules, sometimes associated with an exudate. Microscopic examination of skin nodules reveals the presence of the characteristic encysted spores that contain large cytoplasmic vacuoles. There is no established treatment beyond standard supportive care (Densmore and Green, 2007).

3.5. Parasites Growing concern over amphibian declines and deformities has caused enhanced interest in the effects of parasites on amphibian populations. The parasites can be endoparasites or ectoparasites (Wright, 2006; Densmore and Green, 2007). Parasites affect their hosts in many different ways. In general, heavily parasitized amphibians have a shorter life span, tend to be more susceptible to disease, and have a generally unthrifty appearance (Densmore and Green, 2007). The importance of parasites as a source of mortality and their impact on amphibian population’s dynamics has seldom been addressed. A detailed discussion of amphibian parasites is beyond the scope of this part of dissertation and will be limited to a few cases in which the effect of parasites on the physical condition or survival of amphibians has been

50 INTRODUCTION investigated. Adult clown frogs (Atelopus varius) in Costa Rica are parasitized by larval sarcophagid flies (Notochaeta bufonivora), particularly when the frogs are aggregated near waterfalls (Crump and Pounds, 1985; Pound and Crump, 1987). Fly parasitism could be a major source of mortality in the population, but quantitative data are lacking. Larvae of a tachinid fly, Lucilia bufonivora, sometimes infest population of natterjack toad (Epidalea calamita) and other anurans and can cause substantial mortality of adults (Meisterhans and Heusser, 1970; Sinsch, 1998). Gill (1978) described heavy infestation of leeches in population of red-spotted newt (Notophthalmus viridescens) and speculated that they might be a major factor controlling the population dynamics of newts, either through direct effects of leech attacks or the effects of blood parasites (Trypanosoma) transmitted by the leeches. However, subsequent work showed that trypanosomes had little or no effect on the survivorship of newts or their rates of reproduction (Mock and Gill, 1984; Gill and Mock, 1985); the effects of leech attacks alone were not investigated. Two studies on the effects of infection with parasitic nematode lungworm (Rhabdias) on growth and survival of newly metamorphosed anuran (Bufo bufo and Rana sylvatica) failed to reveal any significant effect (Goater, 1994; Goater and Vandebos, 1997). However, earlier studies with Bufo bufo had shown negative effects of heavier infections by the same parasite on locomotor performance, growth and survival of metamorphs (Goater and Ward, 1992; Goater et al., 1993). Infection by a species of parasitic trematode (Ribeiroia ondatrae) has been implicated in developmental abnormalities seen in several species of anurans, especially the presence of extra limbs (Sessions and Ruth, 1990; Johnson et al., 1999, 2002, 2003; Session et al., 1999; Stopper et al., 2002; Blaustein and Johnson, 2003). Although such deformities have received a lot of news coverage, this is not new phenomenon. There are reports going back nearly 300 years of amphibians with extra limbs (Ouellet, 2000; Johnson et al., 2003). Not all limb malformation can be attribuated to parasite infections; various chemical pollutants may be important as well (Ouellet, 2000; Blaustein and Johnson, 2003). As with other trematodes, fresh water snails (Planorbella) are the intermediate hosts for Ribeiroia, so the prevalence of deformed amphibians may be tightly linked to the population dynamics of the snails. There is evidence that in severely eutrophic ponds impacted by fertilizer runoff or other sorts of pollution, snail population increases, and this may have lead to an increase of amphibians infected with these parasites in recent years (Blaustein and Johnson, 2003; Johnson et al., 2003). The prevalence of deformed individuals is correlated with trematode abundance (Johnson et al., 2001, 2002).

51 INTRODUCTION

Presumably, the fitness consequences for the affected individuals are severe, since their locomotion in impaired and they are likely to be highly susceptible to predation, especially by water birds that are attracted to the awkward movements of deformed frogs. The ecological impact of parasite infections and the resulting limb malformation is not entirely clear. This phenomenon often has been discussed in the context of declining amphibian population (Souder, 2000; Johnson et al., 2002; Blustein and Johnson, 2003). Certainly in some population, deformities in up to 90% of individuals could result in local population extinction, but there is little evidence so far that trematode infections have led to widespread decline of amphibian population (Ouellet, 2000). Indeed, some sites where large numbers of deformed frogs were present more than 50 years ago still support a frog population, with many individulas still infected with parasites (Johnson et al., 2003). Two studies have examined the effects of parasite infection on calling performance and mating success of adult anurans. Hausfater et al. (1990) quantified total numbers of parasitic nematodes and helminthes in various organs of gray tree frogs (Hyla versicolor). Although many individuals were heavily parasitized, there was no relationship between parasite load and measure of performance when males with high and low levels of calling performance were compared for parasite load; there was not a significant difference for any type of parasite. There also was little evidence that parasite infection affected the time of arrival of males at the chorus attendance, or mating success. Tinsley (1990) studied spadefoot toads (Scaphiopus couchii), which usually are heavily infected by the monogenean parasite, Pseudodiplorhis amercanus, which lives in the urinary bladder. Heavily infected toads can be debilitated by depletion of energy reserves, especially during hibernation. Nevertheless, males in a breeding chorus that were captured in amplexus had the same level of parasite infection as those not seen in amplexus. Tinsley (1990) suggested that those males that have been debilitated by parasite infection simply do not come to breeding assemblages at all; whereas males in choruses are not sufficiently impacted by parasites to affect their reproduction success. The effect of parasitism by mites (Hannemania eltoni) on the plethodontid salamander Plethodon angusticlavius was examined in laboratory by Maksimowich and Mathis (2000). Even though heavily infested salamanders were not noticeably unhealthy. Females were tested for the effect of mite infestation on foraging behavior with both fruit flies and termites as prey. Heavily infested females took longer to make a first attack on fruit flies than did females with low parasites loads, but this was not true for females presented with termites. The attack delay averaged only about 75 seconds and was not reflected in lower numbers of

52 INTRODUCTION prey being consumed or in differences in nose-tapping behavior (an indicator of chemosensory exploration). Hence, the ultimate fitness effects of mite infestation are not known, particularly for animals in the field. Nevertheless, in behavioral experiments, there was a tendency for non-parasitized individuals to avoid fecal pellets of heavy parasitized individuals; these pellets are used to mark the territories of individual salamanders (Maksimowich and Mathis, 2001).

3.6. Non-infectious diseases 3.6.1. Heavy metal disorders Heavy metals can be introduced into aqueous environments through industrial and urban effluents, soil leaching, and agricultural runoff as well as through the widespread use of pesticides (McCarthy and Shugart, 1990; Maharajan et al., 2012). These metals can accumulate at higher concentrations in aquatic animals, such as insect larvae, and may affect amphibians in the food chain as a consequence. In captivity, a variety of insect larvae is used as a food source in the amphibian diet. Therefore, a major threat may be associated with the use of dietary food that has been contaminated by the accumulation of heavy metals such as copper, zinc, lead, mercury and cadmium. However, some metals are considered to be trace metals and are essential for normal, biological cell functions. The present section of the dissertation focusses on five heavy metals, including trace metals - also known as trace minerals (copper and zinc) - and xenobiotic or toxic metals (lead, mercury and cadmium) and their possible effects on amphibian health.

3.6.1.1. Trace metals Trace metals occurring at low concentrations (normally < 0.01%) are essential for normal metabolic functions in all living organisms (Halliwell and Gutteridge, 1984; Van Tilborg and Van Assche, 1996). Two distinct functions are performed by essential metals: (1) some of them, such as copper, are involved in electron transfer processes, and (2) the others, such as zinc, mostly participate in governing the reaction mechanisms (Halliwell and Gutteridge, 1984; Cousins and Hempe, 1990; O’Dell, 1990). In general, an element is considered to be an essential micronutrient when: (1) it is consistently determined to be present in all healthy living tissues, (2) their levels are regulated by metabolic processes or homeostasis, and (3) deficiency or defective symptoms are noted due to their depletion or removal from tissue, while those signs can disappear if the elements are returned to the tissue (Wittmann, 1979; Jakimska et al., 2011). Although the specific pathophysiology of metal

53 INTRODUCTION toxicity in amphibians has not been fully described, the basic roles of trace metals seem to be similar in cell biology of the other animals.

3.6.1.1.1. Copper Copper (Cu) contributes to many biochemical activities. It is absorbed from food by the stomach or upper small intestine (duodenum), probably as complexes with amino acids, such as histamine (Halliwell and Gutteridge, 1984). Transfer across the mucosal barrier occurs through non-energy-dependent diffusion (Linder, 1991). However, additional diffusion or carrier-mediated systems in the basolateral membrane play a key role when competition for absorption occurs between copper and zinc (Linder, 1991). Upon entering the blood plasma from the intestinal cell, the copper initially binds to albumin, transcuprein and components of low molecular weight and is transported to the liver where it can be deposited (Linder et al., 1987; Linder, 1991). Subsequently, copper incorporates into the ceruloplasmin in the liver and then transfers into the bloodstream for distribution to different tissues. Although the majority of serum copper is bound to ceruloplasmin (95%) during transport, the rest is bound to albumin, transcuprein, and copper-amino acid complexes. Copper is taken up from the ceruloplasmin via interaction with the cell surface receptors (Linder, 1991). Upon entering the cells, copper plays the role of a cofactor for specific enzymes and electron transport proteins involved in energy or antioxidant metabolism (Halliwell and Gutteridge, 1984; Cousins and Hempe, 1990; O’Dell, 1990). Copper is required for the activity of many enzymes (at least 13) serving as catalytic cofactors in redox chemistry such as: - Ceruloplasmin (antioxidant protection, iron metabolism by involving in Ferroxidase II, and copper transport) - Copper/Zinc superoxide dismutase (antioxidant protection through free radical detoxification) - Cytochrome c oxidase (energy production by electron transport in the mitochondria) - Lysyl oxidase (collagen and elastin formation) - Tyrosinase (pigmentation through the production of melanin) - Metallothionein (copper sequestration) Most features of severe copper deficiency can be explained by a failure of one or more of these copper-dependent enzymes (Halliwell and Gutteridge, 1984; Cousins and Hempe, 1990; O’Dell, 1990; Peña et al., 1999). The adverse effect of low copper intake on reproduction and development were mostly evaluated using a Xenopus embryo-larval development model. A copper-deficient diet under low-copper culture conditions had a significant effect on reproduction and early embryo development, resulting in limb

54 INTRODUCTION teratogenesis (Fort et al., 2000 a, b). Maldevelopment of the hind limbs in particular was found in the F1 generation (Fort et al., 2000). In addition, maldevelopment of the heart, eyes, craniofacial region, brain, and notochord were also observed (Fort et al., 2000 a,b). Although copper is a known essential trace metal, it can be extremely toxic at supraphysiological levels because it produces highly reactive oxygen species (ROS), including hydroxyl radicals (Halliwell and Gutteridge, 1984). These hydroxyl radicals can devastate cellular functions through direct oxidation of proteins, lipid peroxidation in membranes, and cleavage of DNA and RNA molecules (Halliwell and Gutteridge, 1984). Although few research studies have examined the impact of copper on amphibians, the existing studies show that copper overloading in amphibians has negative effects on survivorship (resulting in mortality), swimming performance, percentage of successful metamorphoses (causing delayed metamorphosis) (Herkovits and Helguero, 1998; Parris and Baud, 2004; Chen et al., 2007; Xia et al., 2012). In addition, it causes a reduction in body size and embryonic deformities (Haywood et al., 2004; Chen et al., 2007; Lance et al., 2012). The major excretory route appears to be through the bile, whereas the minimum loss of copper through the urine indicates that any low-molecular-weight copper complexes filtered by the glomeruli are specifically reabsorbed (Linder, 1991; Vallee and Falchuk, 1993).

3.6.1.1.2. Zinc Zinc (Zn) is the second most abundant transition metal ion, after the iron ion (Vasak and Hasler, 2000). Zinc is primarily absorbed by the small intestine (60% in the duodenum, 30% in the ileum, 8% in the jejunum, and 3% through the colon and cecum) and the presence of glucose can assist in its uptake (Davies, 1980; Lee et al., 1989). It is transported from the mucosal barriers into the bloodstream through both passive diffusion and specific zinc transporters. A family of trans-membrane proteins called the Zinc-Iron-related transporter proteins (ZIP) are located in the plasma membrane and facilitate the absorption of zinc into the cells (Tacnet et al., 1990; Gaither and Eide, 2001). Once in a cell, zinc ions move through the membrane channels among various organelles (Yu et al., 2011). In contrast to copper, zinc does not participate in redox reactions; rather, it functions as a Lewis acid to accept a pair of electrons (Williams, 1987). Zinc plays four major biological roles in the organism: - Catalytic role: Zinc is involved as a cofactor in over 300 enzymes, which represent approximately 10% of all proteins that may bind with zinc, such as carbonic anhydrase, alcohol dehydrogenase and alkaline phosphatase (Vallee and Falchuk, 1993; Andreini et al., 2006). These enzymatic functions of zinc control many cell processes, resulting in normal

55 INTRODUCTION

growth, brain development, behavioural responses, reproduction, membrane stability, bone formation and wound healing (Barceloux, 1999; Mocchegiani et al., 2000; Chasapis et al., 2012). - Structural role: Zinc plays structural and functional roles in different proteins involved in DNA replication, reverse transcription, and metallothionein (MT) protein (Mocchegiani et al., 2000; Beyersmann, 2002; Tapiero and Tew, 2003; Chasapis et al., 2012). - Regulatory role: Zinc regulates both the enzymatic levels and the stability of the protein as an activator or an inhibitor ion (Chasapis et al., 2012). To regulate the availability of zinc, eukaryotes first compartmentalise zinc while simultaneously controlling the cellular zinc with the metallothionein/thionein pair (Maret, 2003; Chasapis et al., 2012). - Messenger/Signalling role: Zinc can modulate cellular signal transduction processes such as the modulation of synaptic neurotransmissions in the neurons containing zinc (Mocchegiani et al., 2000; Beyersmann, 2002). In addition, zinc functions as ionic signalling molecules, similar to calcium ions (Yu et al., 2011). For instance, the induction of T-cell proliferative responses is facilitated by increasing extracellular zinc (Yu et al., 2011). Thus, zinc is essential to normal immune functioning (Chasapis et al., 2012). The biological functions of zinc-binding proteins is controlled in different ways at cellular levels, including through zinc transporters and channels, zinc-sensing molecules such as metallothioneins , and the metal-responsive-element-binding transcription factor (Lichtlen and Schaffner, 2001; Eide, 2004). The wide metabolic range governed by zinc refers to the importance of zinc homeostasis in protecting the cell from a deficiency of the metal and from its excess (Bertholf, 1988). The excess zinc compounds are primarily eliminated through excretion, mostly in the faeces and to lesser extendin the urine (Davies and Nightingale, 1975; Wastney et al., 1986). Zinc-deficient cells fail to divide and differentiate, which results in growth impairment, particularly in juvenile cells with high turnover such as skin and gonads cells (Follis et al., 1941; Tucker and Salmon, 1955; Forbes, 1984; Luecke, 1984; Forbes and Yohe, 1960). Zinc deficiency can result in maldevelopment of the mouth, similar to cleft palate, pericardial and ophthalmic edema and axial flexure of the notochord (Fort et al., 1999). Experiments in zinc deficiencies in toad embryos resulted in adults with abnormal ovarian functions and embryos with a high incidence of congenital malformations (Barbieri and De Legname, 1966). Although there is a paucity of data regarding zinc-deficiency symptoms in amphibians compared to fish, the symptoms in amphibians are thought to be similar to those in fish, including increased mortality, reduced growth and appetite, depressed bone calcium and zinc,

56 INTRODUCTION skin erosion, elevated iron and copper concentrations in tissues in the intestine and hepatopancreas, and short-body dwarfism (Ogino and Yang, 1978, 1979; Gatlin and Wilson, 1983; Satoh et al., 1987; Wekell et al., 1983). In contrast to copper, zinc is relatively nontoxic. However, toxicity symptoms can occur with extremely high zinc intake (Bertholf, 1988; Fosmire, 1990). The exact mechanism of intoxication is not fully understood. It seems that zinc toxicity involves the generation of reactive oxygen species, resulting in cell damage and interfering with the metabolism of other ions such as copper, calcium and iron (Peterson, 2001; Daniels et al., 2004). In amphibians, high concentrations of zinc cause different symptoms to appear, including lethargy, poor appetite, darkening skin, dead or deformed embryos, midgut malformations, severe edema of the pericardium and eyes, gut miscoiling, and malformations of the head and mouth. At high, sub-lethal concentrations, severe skeletal kinking, microphthalmia and microencephaly have also been observed (Taban et al., 1982; Woodall et al., 1988; Dawson et al., 1988; Fort et al., 1989; Buhl and Hamilton, 1990). Zinc has a toxic effect on larval gonads, especially on germ cells in the ovarian structure (Gipouloux, 1986). In addition, zinc overloading increases the time to complete metamorphosis and decreases mass at time of metamorphosis or reduces body size (Bishop et al., 2000; Brodeur et al., 2009; Camponelli et al., 2009). Christensen et al. (2004) showed that increasing zinc concentrations caused a significant decrease in sperm motility percentages due to its interference with cellular processes (e.g. cellular respiration, flagellar bending and ion exchange), which thereby inhibited sperm motility.

3.6.1.2. Xenobiotic metals Xenobiotic metals (Xenos originates from the Greek for “foreign”, and bios means “life”), or toxic metals, lack known biological functions. They can attach to enzymes and cellular receptors instead of essential minerals (Singh, 2007). They compromise or block the metabolic processes, depending on that particular enzyme or cellular receptor, and effectively reduce the uptake of needed minerals by the body’s cells and enzymes, which leads to disturbance of growth, reproduction, the immune system, and the metabolic functions (Hanas and Gunn, 1996; Singh, 2007; Jakimska et al., 2011).

3.6.1.2.1. Lead Lead (Pb) is acutely toxic (genotoxic and neurotoxic) to amphibians and accumulates in significant amounts in the liver from the lowest exposure concentration (Eisler, 1988; Mouchet et al., 2007). The toxicokinetic effects of lead include substitution of and

57 INTRODUCTION competition with Ca2+, disruption of Ca2+ homeostasis, binding with sulfhydryl groups, stimulating the release of Ca2+ from mitochondria, damaging mitochondria and mitochondrial membranes, replacing zinc in zinc-mediated processes, increasing oxidative stress, inhibiting anti-oxidative enzymes and altering lipid metabolism (Needleman, 1991; Ahameda and Siddiqui, 2007). Chen et al. (2006) showed that lead overloading resulted in abnormal swimming behaviours, which was associated with spinal deformities in amphibians. In addition, the maximum swimming speed and growth rate of tadpoles were significantly decreased. Lead exposure causes different symptoms to appear in amphibians, including sloughing off of the skin, hypertrophy of the liver, spleen and stomach, decreased muscle tone, excessive bile secretion, haematological effects (decreasing erythrocytes, neutrophils and monocytes), salivation, muscular twitching, and delayed metamorphosis (Eisler, 2000; Sparling et al., 2000; Ezemonye and Enuneku, 2005; Bazar et al., 2010). Lead is positively associated with a significant decrease in the activity of the delta-aminolevulinic acid dehydratase (delta-ALAD activity) and an increase in the free erythrocyte protoporphyrin (FEP) levels, which results in impaired heme synthesis (Arrieta et al., 2004). Lead can negatively affect the functioning of the polymorphonuclear cells, thereby decreasing the phagocytic and lytic activity (Fink and Salibian, 2005). In addition, it causes the production of natural antibodies to significantly increase. Thus, lead overloading may act as an immunostimulating factor on amphibian hormonal immune systems (Fink and Salibian, 2005).

3.6.1.2.2. Mercury Mercury (Hg) is highly toxic in its organic form, methylmercury (MeHg), occurring in the organism (Watras and Bloom, 1992; Hill et al., 1996). The mechanisms of action include protein precipitation and enzyme inhibition through binding to sulfhydryl, phosphoryl, carboxyl, amide, and amine groups. In addition, it can cause biochemical damage by interrupting intracellular calcium homeostasis, mitochondrial damage, lipid peroxidation, microtube destruction and the neurotoxic accumulation of serotonin, aspartate and glutamate (Yee and Choi, 1996). There is limited data available on mercury toxicity in amphibians. However, the adverse effects of mercury exposure reported commonly among all studies include its neurotoxicity, immunotoxicity (immunosuppression), endocrine disruption, impaired reproduction, physical

58 INTRODUCTION malformation, and mortality in fish and amphibians (Birge et al., 1979; Weiner and Spry, 1996; Wolfe et al., 1998; Eisler, 2006; Scheuhammer et al., 2007; Tan et al., 2009; Wada et al., 2009; Grillitsch and Schiesari, 2010). Since mercury can degenerate the neuron function, resulting in disruption of the brain’s ability to effectively control motor functions, the behavioural disorders may inhibit the ability of amphibians to successfully execute tasks critical to survival (e.g. reducing motivation to capture prey, avoiding predators, or successfully competing with others) (Little et al., 1990; Walker et al., 2005; Burke et al., 2010). Todd et al. (2012) recently reported that adverse effects of maternal mercury exposure in larval amphibians might persist to affect later terrestrial life stages.

3.6.1.2.3. Cadmium Cadmium (Cd) is a toxic and potentially genotoxic substance for amphibians (Mouchet et al., 2007). Amphibians can be affected by very low amounts of cadmium and it especially accumulates in the cytosolic fraction of the liver and kidneys (Zettergren et al., 1991; Vogiatzis and Loumbourdis, 1997). At the molecular level, the mechanisms of cadmium toxicity include interference with or substitution of essential metals particularly for Ca2+or Zn2+, disruption of signalling and biomolecular functions by binding to the sulphur (-SH) groups of proteins, cysteine, and glutathione (Brzóska and Moniuszko-Jakoniuk, 1998, 2001; Bertin and Averback, 2006). Consequently, the production of reactive oxygen species (ROS) increases the changes in the expression of different genes, resulting in cell cycle arrest or apoptosis (Min et al., 2000; Chan et al., 2006). In addition, cadmium can directly affect the gene expression through the inhibition of DNA methylation and it acts as an epigenetic or indirect genotoxic carcinogen (Waalkes and Misra, 1996; Takiguchi et al., 2003). Amphibians exposed to cadmium display a variety of symptoms, including malformation, delayed and arrested development indicating a major inhibitory effect on metamorphosis, decreasing oogenesis, decreased larval weight due to loss of appetite, and decreased survival percentages (Calevro et al., 1998; Lienesch et al., 2000; Flament et al., 2003; James and Little, 2003; Gross et al., 2007). In addition, cadmium can affect vulnerability (e.g. predation and fertilisation) in sub-lethal concentrations, while it causes immediate death in lethal concentrations (Mouchet et al., 2007; Gross et al., 2007). Contaminated amphibians exhibit high proportions of B-cells expressing immunoglobulin (Ig)M in mesonephros and in the liver, suggesting that cadmium ions act directly on B-cells, perhaps at the membrane level (Zettergren et al., 1991). However, cadmium inhibits early events in T-cell functions by interfering with signal transduction and T- cell activation

59 INTRODUCTION

(Cifone et al., 1989). Therefore, cadmium may affect antibody and Ig synthesis indirectly in amphibian through its interaction with regulatory T-cells (Zettergren et al., 1991).

60 INTRODUCTION

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SCIENTIFIC AIMS

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Scientific Aims of the Study

Amphibian populations are at the forefront of the global biodiversity crisis, with more extinct or declining species than any other class of vertebrates. Of the fifteen Belgian amphibian species, 10 are decreasing in population and currently threatened. Although putative causes of observed declines are numerous and frequently controversial, the emergence of infectious diseases such as chytridiomycosis and ranavirosis, and invasive species such as American bullfrogs (Lithobates catesbeianus), represent two of the greatest threats confronting native amphibians. One of the main issues hampering amphibian conservation is the limited knowledge with regard to amphibian infections, which is currently heavily focused on chytridiomycosis and ranavirosis. The general aim of this PhD study was, therefore, to gain insight in microbiological infections in native amphibians, which are of importance in amphibian conservation related issues. The first specific aim was to determine the presence of bacteria, viruses and fungi in native and invasive amphibians with the potential to affect amphibian health. For this purpose, 1) we determined the prevalence of Batrachochytrium dendrobatidis (causative agent of chytridiomycosis) and ranaviruses in native populations of common toads (Bufo bufo); 2) we investigated the presence of ranaviruses, B. dendrobatidis and Chlamydiales in invasive American bullfrogs (L. catesbeianus). The second aim was to estimate the risk of live feed items to the health of indigenous amphibian communities and to ex situ amphibian conservation. A typical amphibian feed item are bloodworms (aquatic larvae of Chironomidae). These bloodworms either are collected in Belgium or abroad and large quantities are used for feeding aquatic animals and for recreational fishing. As such, they may pose a risk of vectoring pathogens or toxins such as heavy metals into indigenous amphibian communities or in ex situ amphibian conservation programmes. The third specific aim was to determine the zoonotic risk of human contact with amphibians during conservation activities such as safeguarding activities for native amphibians and controlling invasive amphibian species. For this purpose, 1) we examined the presence of Salmonella sp. in native common toads (Bufo Bufo); 2) we determined the occurrence of zoonotic agents, i.e. Coxiella burnetii, Neospora caninum, Leptospira sp., Toxoplasma gondii, Mycoplasma sp., Campylobacter sp., Salmonella sp. and broad-spectrum beta-lactamase producing Escherichia coli in invasive American bullfrogs in Belgium.

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Road-killed common toads (Bufo bufo) in Flanders (Belgium) reveal low prevalence of ranaviruses and Batrachochytrium dendrobatidis

An Martel,1,5 Mojdeh Sharifian Fard,1,5 Pascale Van Rooij,1 Robert Jooris,2 Francine Boone,3 Freddy Haesebrouck,1 David Van Rooij,4 and Frank Pasmans1

1. Department of Pathology, Bacteriology and Avian Diseases, Ghent University Salisburylaan 133, B-9820 Merelbeke, Belgium 2. Natuurpunt–Hyla Coxiestraat 11, 2800 Mechelen, Belgium 3. Nachtegaalstraat 91, 1501 Buizingen, Belgium 4. Renard Centre of Marine Geology, Department of Geology and Soil Science, Ghent University, Krijgslaan 281 S8, B-9000 Ghent, Belgium 5. These authors contributed equally to this work

Adapted from Journal of Wildlife Diseases, 2012, 48(3): 835–839

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Abstract Road-killed common toads (Bufo bufo; n=1,740) from Flanders, Belgium, were tested for ranaviruses and Batrachochytrium dendrobatidis using polymerase chain reaction. Both infections were present at a very low prevalence (<0.2% with a confidence interval of 95%) for ranaviruses and 0.63% for B. dendrobatidis.

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Healthy amphibians can be asymptomatic carriers of diseases responsible for devastating amphibian population declines. Most importantly, ranaviruses and B. dendrobatidis are primary pathogens of amphibians, and disease outbreaks are believed to be driven by environmental factors (Walker et al., 2010). A thorough risk analysis to predict future scenarios in indigenous amphibian species should include the identification of principle pathogen reservoirs. The difficulty in identifying amphibian, clinically healthy reservoirs for both agents is that asymptomatically infected animals often have low-grade infections. Although noninvasive sampling (e.g., skin swabs) enables the detection of both pathogens, tissue samples provide a higher detection probability. However, conservation concerns hamper tissue sampling of wild amphibians. Traffic kills many amphibians on roads yearly, especially during spring migration (Elzanowski et al., 2009). These road victims could provide a good opportunity for monitoring ranaviruses and B. dendrobatidis in native amphibians. In Flanders (Belgium), the common toad (Bufo bufo) provides the most obvious choice to monitor both infections for four reasons: Common toads are commonly killed by traffic during spring migration; migrating toads are counted on numerous sites in Flanders, enabling an estimate of population trends; the common toad is a suitable host for both ranaviruses and B. dendrobatidis (Bosch and Marinez-Solano, 2006; Cunningham et al., 2007a,b); and toad populations have been found to be declining in several regions (Loras et al., 2011). Our goal was to assess ranaviruses and B. dendrobatidis infections in common toads using tissues of road-killed specimens during the migration season. Between 27 February 2011 and 7 April 2011, 1,740 road-killed common toads were collected from 104 migration sites in all five Flemish provinces (Figure. 1). Each toad was placed in a plastic bag that was sealed and labeled. For detection of B. dendrobatidis DNA, a quantitative polymerase chain reaction (qPCR; Boyle et al., 2004) was performed on a skin sample of each toad. For detection of ranavirus infections, a liver sample was examined by PCR using the primers MCP 4 and MCP 5 (Mao et al., 1997). To assess whether the presence of one or both pathogens in a toad population coincides with obvious population declines, if available, numbers of migratory toads of 2011 and previous years were extracted using the database obtained from the working group Natuurpunt Hyla (Hyla Werkgroep, 2012). None of the samples were PCR-positive for ranaviruses. At nine sites (three in the province of East Flanders, three in Flemish Brabant, and three in Limburg), 11 B. dendrobatidis-positive toads were found (0.63% prevalence; Figure 1).

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Figure 1. Locations in Flanders (Belgium) where road-killed common toads (Bufo bufo) were collected during the migration season from 27 February 2011 to 7 April 2011. Locations where B. dendrobatidis was found are indicated by black circles and negative locations by grey circles. FR=France; NL=The Netherlands; D=Germany; W.-Fl.=West-Flanders; E.-Fl.=East Flanders; Antw.=Antwerp; Fl.- Br.=Flemish Brabant; B.=Brussels; Lim.=Limburg; R.=river.

Levels of B. dendrobatidis in the positive samples were relatively low, with an average ±SD of 0.25±60.11 genomic equivalents. Data on the number of migrating toads were available for four of the nine infected sites. For three additional sites, numbers of migrating toads were available within a region of 5 km. For all positive sites, no significant decrease in the number of migrating toads was observed during the years monitored (Table 1). This study demonstrates that if ranavirus infections are present in adult common toads in Flanders, the prevalence is very low (< 0.2% with a confidence interval of 95%). Thus, we can reject the hypothesis that, at present, adult common toads provide an important reservoir for ranaviruses in the area examined. However, these results should not be overinterpreted. First, only adult toads were sampled during the migration season, implying that these toads survived hibernation well and were healthy enough to show reproductive (migratory) behavior. Second, all samples were collected during the relatively short migration period and thus might not be representative of the entire population. Third, it is possible that the adults survived a past ranavirus infection and have developed antibodies, as was shown for Bufo marinus (Zupanovic et al., 1998).

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Table 1. Yearly numbers of mugrating common toads (Bufo bufo) at a B. dendrobatidis- positive site in Flanders (Belgium). Data from Natuurpunt Hyla (Hyla Werkgroep. 2012)a

No. positive Site (Province) GPS coordinates samples/no. 2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 tested N 51°14'38.30" Beveren (East-Flanders)a 2/12 ND 609 655 852 1204 855 1038 817 696 1174 E 4°17'39.03"

N 50°57’03.98” Munte (East-Flanders) 1/48 986 799 1253 1603 1712 1947 937 819 1129 679 E 3°45’00.00”

N 50°49’28.73” Ninove (East-Flanders)a 1/70 ND ND ND ND ND ND ND 391 524 ND 3°58’53.58”

N 50°43’19.03” Dworp (Flemish Brabant)a 2/144 ND ND ND ND 761 731 ND 663 1995 3651 E4°17’31.77”

N 50°52’53.76” Diepenbeek (Limburg) a 1/1 ND ND ND 1027 1322 713 1060 495 ND ND E 5°24’48.86”

N 51°00’47.76” Dilsen-stokkem (Limburg) b 1/5 3594 3886 3577 4062 3993 2073 1809 1077 1782 2244 E5°39’03.04”

N 50°55’52.72” Hasselt (Limburg) a, b 1/79 ND ND 729 708 ND 211 134 ND 168 334 E 5°19’08.83”

N50°42’31.96” Bever (Flemish Brabant)a 1/35 ND ND ND ND ND ND ND ND ND ND E 3°58’25.96”

Sint Pieters Kapelle (Flemish N 50°41’49.95” 1/90 ND ND ND ND ND ND ND ND ND ND a Brabant) E3°57’59.56” a. ND=no data available b. Data recorded at a location within 5 km of the positive sample

Our data suggest that ranaviruses might be absent from some native common toad populations. If true, then introduction of a ranavirus into these populations can result in disease outbreaks. Furthermore, introduction of a novel ranavirus into any population can result in mass morbidity or mortality. Neither scenario is unlikely because ranaviruses have been demonstrated in captive amphibians in Flanders (Pasmans et al., 2008), in exotic invasive bullfrogs (Lithobates catesbeianus) in Flanders (Sharifian Fard et al., 2011), and recently in a large ranavirus outbreak in the neighboring Netherlands (Kik et al., 2011). The low prevalence of B. dendrobatidis in the study area stresses the importance of a large sample size if the presence of B. dendrobatidis in a seemingly healthy amphibian population needs to be excluded. Population trends in infected common toad populations, as derived from data on the migrating toads, are not suggestive of population declines over the last 5 years. Moreover, the authors of this paper are the referral center for disease outbreaks of

99 STUDY 1 amphibians in Flanders, and no proven outbreak of chytridiomycosis has yet been confirmed. The only confirmed case of lethal chytridiomycosis was an isolated case in a midwife toad (Alytes obstetricans) in the Belgian Ardennes (Pasmans et al., 2010). The combination of low prevalence, low B. dendrobatidis load in infected animals, absence of obvious negative population trends, and absence of confirmed chytridiomycosis outbreaks, suggests that B. dendrobatidis is endemic at a low intensity in the Flemish region. However, the pathogenic potential of B. dendrobatidis has been demonstrated on numerous occasions, extirpating entire amphibian species (Stuart et al., 2004; Wake and Vredenburg, 2008). Again, our results should be interpreted carefully. Besides the caveats mentioned above, seasonality of B. dendrobatidis infection dynamics is well documented. The occurrence of clinical chytridiomycosis depends on numerous factors, including strain virulence, host susceptibility, and environmental parameters. Given current climate change, epidemics might be expected in the future. Thus, vigilance systems would be advisable to monitor B. dendrobatidis infection dynamics.

Acknowledgment We thank Natuurpunt volunteers who sampled road victims.

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References Bosch J, Martinez-Solano I. 2006. Chytrid fungus infection related to unusual mortalities of Salamandra salamandra and Bufo bufo in the Penalara natural park, Spain. Oryx 40:84–9. Boyle DG, Boyle DB, Olsen V, Morgan JAT, Hyatt AD. 2004. Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real- time taqman PCR assay. Diseases of Aquatic Organisms 60:133–9. Cunningham AA, Hyatt AD, Russel P, Bennett PM. 2007. Experimental transmission of a Ranavirus disease of common toads (Bufo bufo) to common frogs (Rana Temporaria). Epidemiology and Infection 135:1213–6. Elzanowski A, Ciesiolkiewicz J, Kaczor M, Radwariska J, Urabn R. 2009. Amphibian road mortality in Europe: a meta-analysis with new data from Poland. European Journal of Wildlife Research 55:33–43. Hylawerkgroep. 2012. Natuurpunt Hyla. Available at http://www.Hylawerkgroep.Be/. Accessed March 2012. Kik M, Martel A, Spitzen-Van Der Sluijs A, Pasmans F, Wohlsein P, Gröne A, Rijks JM. 2011. Ranavirus-associated mass mortality in wild amphibians, the Netherlands, 2010. A first report. Veterinary Journal 190:284–6. Loras AF, Hidalgo-Vila J, Hermosilla C, Garcia G, Lopez J, Duffus ALJ, Cunningham AA, Raco V. 2011. Preliminary health screening and possible pathogen determination in a Bufo bufo (Linnaeus, 1758) (Amphibia: Bufonidae) population. Journal Of Natural History 45:1–14. Mao J, Hedrick RP, Chichar VB. 1997. Molecular characterization, sequence analysis, and taxonomic position of newly isolated fish Iridoviruses. Virology 229:212–20. Pasmans F, Blahak S, Martel A, Pantchev N, Zwart P. 2008. Ranavirus-associated mass mortality in imported red tailed knobby newts (Tylototriton kweichowensis): a case report. Veterinary Journal 176:257–9. Pasmans F, Muijsers M, Maes S, Van Rooij P, Brutyn M, Ducatelle R, Haesebrouck F, Martel A. 2010. Chytridiomycosis related mortality in a midwife toad (Alytes obstetricans) in Belgium. Flemish Veterinary Journal 79:460–2. Sharifian Fard M, Pasmans F, Adriaensen C, Devisscher S, Adriaens T, Louette G, Martel A. 2011. Ranavirosis in invasive bullfrogs, Belgium. Emerging Infectious Diseases 17: 2371–2. Stuart SN, Chanson JS, Cox NA, Young BE, Rodrigues ASL, Fischman DL, Waller RW.

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2004. Status and trends of amphibian declines and extinctions worldwide. Science 306:1783–6. Wake DB, Vredenburg VT. 2008. Are we in the midst of the sixth mass extinction? A view from the world of amphibians. Proceedings of the National Academy of Sciences USA 105: 11466–73.

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STUDY 2

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104 STUDY 2

Ranavirosis in invasive bullfrogs, Belgium

Mojdeh Sharifian-Fard1, Frank Pasmans1, Connie Adriaensen1, Sander Devisscher2, Tim Adriaens2, Gerald Louette2, and An Martel1

1. Department of Pathology, Bacteriology and Avian Diseases, Ghent University Salisburylaan 133, B-9820 Merelbeke, Belgium 2. Research Institute for Nature and Forest, Kliniekstraat 25, B-1070 Brussels, Belgium

Adapted from Emerging Infectious Diseases, 2011, 17(12): 2371-2

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Ranavirosis is caused by icosahedral cytoplasmic DNA viruses that belong to the family Iridoviridae, in particular by 4 species of Ranavirus: Frog Virus 3 (FV3), Bohle iridovirus, Ambystoma tigrinum virus, and a possible species Rana catesbeianus virus Z. In Europe, FV3 has been identified in several outbreaks of ranavirosis, characterized by mass deaths, notably in green frogs (Pelophylax sp.) in Denmark, Croatia, and the Netherlands (Fijan et al., 1991; Ariel et al., 2009); Rana temporaria and Bufo bufo in the United Kingdom (Cunningham et al., 1996; Hyatt et al., 2000); and Alytes obstetricans and Ichthyosaura alpestris in Spain (Balseiro et al., 2009). The invasive exotic bullfrog (Lithobates catesbeianus) has been introduced in several European countries and has established large breeding populations in France, Italy, Germany, Greece, and Belgium (Ficetola et al., 2007). In addition to their direct effect on native amphibians through competition and predation, bullfrogs are thought to be carriers of chytridiomycosis (Schloegel et al., 2010; Daszak et al., 2004) and, possibly, ranaviruses. Although mass deaths of L. catesbeianus tadpoles has been reported in aquaculture facilities, L. catesbeianus tadpoles are generally considered a subclinical reservoir of ranaviruses in the United States (Miller et al., 2009). To assess the role of bullfrogs as carriers of ranaviruses in Europe, we collected 400 clinically healthy tadpoles of L. catesbeianus from 3 invasive bullfrog populations at Hoogstraten, Belgium (51°47′Ν, 4°75′Ε) during May–June 2010. All larvae were euthanized as part of an invasive species eradication project and stored at –20°C until further use. At necropsy, liver tissues were collected, and DNA was extracted by using the Genomic DNA Mini Kit (BIOLINE, London, UK). PCR to detect ranaviruses was performed as described by Mao et al. (Mao et al., 1997). Three samples showed positive results with this PCR. These samples were sequenced by using primers M4 and M5 described by Mao et al., (1997) and blasted in GenBank. A 100% homology with the common midwife toad (A. obstetricans) ranavirus partial major capsid protein gene (GenBank accession no. FM213466.1) was found (Balseiro et al., 2009). Despite the low prevalence of ranavirus infection (0.75%) in the bullfrog tadpoles examined, this study shows that invasive bullfrogs, a known reservoir of chytridiomycosis, are also a likely carrier of ranaviral disease in Europe.

Acknowledgment This study was partly performed in the framework of European Union Interreg IVA project IVA-VLANED-2.31 “Invasieve exoten in laanderen en Zuid-Nederland–IN EXO.”

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References Ariel E, Kielgast J, Svart HE, Larsen K, Tapiovaara H, Jensen BB. 2009. Ranavirus in wild edible frogs Pelophylax kl. esculentus in Denmark. Diseases of Aquatic Organisms 85:7–14. Fijan N, Matasin Z, Petrinec Z, Valpotiç I, Zwillenberg LO. 1991. Isolation of an Iridovirus- like agent from the green frog (Rana esculenta L.). Vet Zagreb. 3:151–8. Cunningham AA, Langton TES, Bennet PM, Lewin JF, Drury SEN, Gough RE. 1996. Pathological and microbiological findings from incidents of unusual mortality of the common frog (Rana temporaria). Philosophical Transactions of the Royal Society B: Biological Sciences 351:1539–57. Hyatt AD, Gould AR, Zupanovic Z, Cunningham AA, Hengstberger S, Whittington RJ. 2000. Comparative studies of piscine and amphibian Iridoviruses. Archives of Virology 145:301–31. Balseiro A, Dalton KP, Del Cerro A, Marquez I, Cunningham AA, Parra F, Pathology. 2009. Isolation and molecular characterisation of a Ranavirus from the common midwife toad Alytes obstetricans on the Iberian Peninsula. Diseases of Aquatic Organisms 84:95–104. Ficetola GF, Coic C, Detaint M, Berroneau M, Lorvelec O, Miaud C. 2007. Pattern of distribution of the American bullfrog Lithobates catesbeianus in Europe. Biological Invasions 9:767–72. Schloegel LM, Ferreira CM, James TY, Hipolito M, Longcore JE, Hyatt AD. 2010. The North American bullfrog as a reservoir for the spread of Batrachochytrium dendrobatidis in Brazil. Animal Conservation 13(S1):53–61. Daszak P, Strieby A, Cunningham AA, Longcore JE, Brown CC, Porter D. 2004. Experimental evidence that the bullfrog (Lithobates catesbeianus) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians. Journal of Herpetology 14:201–7. Miller DL, Gray MJ, Rajeev S, Schmutzer AC, Burton EC, Merril A. 2009. Pathological findings in larval and juvenile anurans inhabiting farm ponds in Tennessee, USA. Journal of Wildlife Diseases 45:314–24. Mao J, Hedrick RP, Chichar VB. 1997. Molecular characterization, sequence analysis, and taxonomic position of newly isolated fish iridoviruses. Virology 229:212–20.

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The novel ‘Candidatus Amphibiichlamydia ranarum’ is highly prevalent in invasive exotic bullfrogs (Lithobates catesbeianus)

An Martel,1 Connie Adriaensen,1 Mojdeh Sharifian-Fard,1 Mado Vandewoestyne,2 Dieter Deforce,2 Herman Favoreel,1 Karolien Bergen,1 Annemarieke Spitzen-van der Sluijs,4 Sander Devisscher,3 Tim Adriaens,3 Gerald Louette,3 Kristof Baert,3 Alex Hyatt,5 Sandra Crameri,5 Freddy Haesebrouck1 and Frank Pasmans1

1. Department of Pathology, Bacteriology and Avian Diseases, Ghent University Salisburylaan 133, B-9820 Merelbeke, Belgium 2. Laboratory for Pharmaceutical Biotechnology, Faculty of Pharmaceutical Sciences, Ghent University, Harelbekestraat 72, B-9000 Ghent, Belgium 3. Research Institute for Nature and Forest, Kliniekstraat 25, B-1070 Brussels, Belgium 4. RAVON, Postbus 1413, 6501 BK Nijmegen, The Netherlands 5. Australian Animal Health Laboratory, Private Bag 24, VIC 3219 East Geelong, Australia

.

Adapted from Environmental Microbiology Reports, 2013, 5(1): 105–108

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Abstract Knowledge concerning microbial infectious diseases in the current amphibian crisis is rudimentary and largely limited to ranavirosis and chytridiomycosis. The family Chlamydiaceae is gaining attention as a common cause of disease in amphibians and may harbour new and emerging amphibian pathogens. We identified a novel species of Chlamydiales (Candidatus Amphibiichlamydia ranarum) with a prevalence of 71% in exotic invasive bullfrog tadpoles (Lithobates catesbeianus) from an introduced population in the Netherlands. The sequence of a 1474 bp 16S rRNA gene fragment showed that the novel taxon forms a well-defined clade with ‘Candidatus Amphibiichlamydia salamandrae’ within the Chlamydiaceae family. Although none of the tadpoles examined showed signs of clinical disease, urgent evaluation of its pathogenic potential for native amphibian species is required.

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Exotic invasive bullfrogs (Lithobates catesbeianus) are considered to exert a considerable negative impact on native amphibian communities (Lever, 2003; Ficetola et al., 2007). Bullfrogs are a notorious source of the infectious diseases chytridiomycosis and ranavirosis, affecting amphibian populations globally (Garner et al., 2006; Sharifian-Fard et al., 2011). However, little is known regarding the presence of other microbial agents that might be transferred to native amphibians. The family Chlamydiaceae is gaining attention as a common cause of disease in amphibians and is associated with high mortality rates in a variety of amphibian species. Of the order Chlamydiales, the presence of the species Chlamydia psittaci, C. pneumoniae, C. abortus, C. suis and the novel genus ‘Candidatus Amphibiichlamydia salamandrae’ has been demonstrated in amphibians (Newcorner et al., 1982; Wright, 1996; Mutschmann, 1998; Berger et al., 1999; Reed et al., 2000; Hotzel et al., 2001; Blumer et al., 2007; Martel et al., 2012). This new genus may represent a clade within the chlamydiae adapted to amphibians, possibly containing new and emerging amphibian pathogens. In this study, we examined whether and which species of Chlamydiales occur in invasive and reproducing exotic bullfrog populations in Belgium (Hoogstraten, Antwerp) and the Netherlands (Baarloo, Noord Limburg). Using PCR for the detection of the 16S rRNA gene of Chlamydiales (Everett et al., 1999), we demonstrated the presence of Chlamydiales DNA in 71% of the 200 livers from Dutch tadpoles examined but in none of those from the 200 Belgian tadpoles. A 1473 bp 16S rRNA gene fragment was sequenced from 10 of these tadpoles using the primers 16SF and 16SR, described by Everett (2000). This sequence was identical for all tadpoles and shared more than 90% nucleotide identity with known members of the order Chlamydiales and 91% identity with the reference 16S rRNA gene sequence of C. abortus B577 (Accession No. D85709). Immunofluorescently stained (IMAGEN Chlamydia kit, Oxoid, Basingstoke, UK) impression smears from tadpole livers revealed the sporadic presence of strongly fluorescent aggregates associated with anuran cells in the liver in 7/20 (35%) of the livers examined, resembling Chlamydiales in host cells (Figure 1). PCR confirmed the presence of Chlamydiales DNA in these livers that showed positive fluorescent staining. An additional 10/20 livers were also PCR positive but did not yield a positive result on immunofluorescent (IF) staining. The three remaining IF negative samples were also PCR negative. Excision of 100 fluorescent particles using laser capture microdissection (Vandewoestyne et al., 2012) with subsequent PCR and sequencing confirmed colocalization of the novel Chlamydiales DNA sequence with fluorescent particles. Despite intensive searching, transmission electron

113 STUDY 3 microscopic (TEM) examination of PCR and IF positive samples did not result in the detection of Chlamydiales like organisms. In amphibians infected with the closely related Candidatus Amphibiichlamydia salamandrae, the presence of Chlamydiales like organisms in the host cells could be clearly demonstrated using TEM and IF revealed the presence of numerous positively stained inclusions in the host cells. These animals exhibited marked clinical signs (Martel et al., 2012). The absence of clinical signs, low numbers of IF positive cells and absence of relevant TEM findings suggest low-level infections in the tadpoles or perhaps even symbiosis in the present study (Horn, 2008). However, the consistent PCR and sequencing findings combined with IF demonstrate the presence of a novel Chlamydiales taxon, despite the lack of relevant morphological data. Based on the sequences obtained, the novel taxon can be identified as a member of the Chlamydiaceae (Everett et al., 1999), with highest 16S rRNA similarity to ‘Candidatus Amphibiichlamydia salamandrae’ (95%). Neighbour-joining analysis (Kodon; Applied Maths, Sint-Martens-Latem, Belgium) showed that, together with ‘Candidatus Amphibiichlamydia salamandrae’, the novel taxon forms a distinct branch in the well-supported monophyletic clade with the genera Chlamydia and Candidatus Clavochlamydia salmonicola (family Chlamydiaceae) (Figure 2).

Figure 1. Immunocytochemically stained (IMAGEN Chlamydia kit, Oxoid, Basingstoke, UK) impression smears from tadpole livers.

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Figure 2. Topology of the novel species ‘Candidatus Amphibiichlamydia salamandrae’ within the phylogenetic tree obtained by neighbour joining and based on 16S rRNA gene data from representative species. Maximum parsimony and UPGMA analyses yielded cladograms with the same topology (results not shown). Numbers show the percentage of times each branch was found in 1000 bootstrap replicates. The tree has been rooted with Verrucomicrobium spinosum as outgroup.

The discovery of this second species suggests the genus Amphibiichlamydia to contain species adapted to amphibians. Further studies should focus on the ability of this genus to cross vertebrate taxon barriers as recently shown for C. pneumoniae (Mitchell et al., 2010). The presence of the novel taxon in the bullfrogs tadpoles was not associated with obvious clinical disease, resembling results obtained by Blumer and colleagues (2007), who found a prevalence of 2.5% of Chlamydiales in 126 Rana temporaria without a clear link with disease. However, its closest relative ‘Candidatus Amphibiichlamydia salamandrae’ has been associated with severe disease and high mortality (Martel et al., 2012). Infection trials using an isolate of the novel taxon should clarify its pathogenic potential to bullfrogs and native amphibian species. However, attempts to isolate this novel species on a variety of cell lines have failed so far. Until the pathogenic potential of these novel Chlamydiales is evaluated, we recommend to include diagnostics for Chlamydiales in any outbreak of amphibian disease, both in captive and in wild populations. Indeed, this study demonstrates our current lack of

115 STUDY 3 knowledge of amphibian pathogens, other than the few well-studied ones such as chytridiomycosis or ranavirosis (Duffus, 2009). ‘Candidatus Amphibiichlamydia ranarum’ [Am.phi. bi.i.chla.my’dia. N.L. n. Amphibia name of host class; L. fem. n. Chlamydia name of bacterial taxon; N. L. fem. n. Amphibiichlamydia Chlamydia from an amphibian; ra.na’rum L. gen. n. from frogs]. The provisional taxon ‘Candidatus Amphibiichlamydia ranarum’ contains cell-associated bacteria that infect frogs of the genus Lithobates in freshwater environments. The 16S rRNA gene of ‘Candidatus Amphibiichlamydia ranarum’ has been deposited in the GenBank with Accession number JN402380 and shows phylogenetic affinity towards the family Chlamydiaceae.

Acknowledgements This study was partly performed in the framework of EU Interreg IVA project IVA- VLANED-2.31 ‘Invasieve exoten in laanderen en Zuid-Nederland – IN EXO’.We are grateful to Jean Euzéby for support with the nomenclature.

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References Berger L, Volp K, Mathews S, Speare R, Timms P. 1999. Chlamydia pneumoniae in a free- ranging giant barred frog (Mixophyes iterates) from Australia. Journal of Clinical Microbiology 37: 2378–2380. Blumer C, Zimmermand DR,Weilenmann R, Vaughan L, Pospischil A. 2007. Chlamydiae in free-ranging and captive frogs in Switzerland. Veterinary Pathology 44: 144–150. Duffus ALJ. 2009. Chytrid blinders: what other disease risks to amphibians are we missing? Ecohealth 6: 335–339. Everett KDE. 2000. Chlamydia and Chlamydiales: more than meets the eye. Veterinary Microbiology 75: 109–126. Everett KDE, Bush RM, Andersen AA. 1999. Emended description of the order Chlamydiales, proposal of Parachlamydiaceae fam. nov. and Simkaniaceae fam. nov., each containing one monotypic genus, revised taxonomy of the family Chlamydiaceae, including a new genus and five new species, and standards for the identification of organisms. International Journal of Systematic Bacteriology 49: 415–440. Ficetola GF, Thuiller W, Miaud C. 2007. Prediction and validation of the potential global distribution of a problematic alien invasive species – the American bullfrog. Diversity and Distributions 13: 476–485. Garner TWJ, Perkins MW, Govindarajulu P, Seglie D, Walker S, Cunningham AA, Fisher, MC. 2006. The emerging amphibian pathogen Batrachochytrium dendrobatidis globally infects introduced populations of the North American bullfrog, Rana catesbeiana. Biology Letters 2:455–459. Horn M. 2008. Chlamydiae as symbionts in eukaryotes. Annual Review of Microbiology 62: 113–131. Hotzel H, Grossman E, Mutschmann F, Sachse K. 2001. Genetic characterization of a Chlamydophila pneumoniae isolate from an African frog and comparison to currently accepted biovars. Systematic and Applied Microbiology 24: 63–66. Lever C. 2003. Naturalized Amphibians and Reptiles of the World. New York, USA: Oxford University Press. Martel A, Adriaensen C, Bogaerts S, Ducatelle R, Favoreel H, Crameri S, et al. 2012. Novel Chlamydiaceae disease in captive salamanders. Emerging Infectious Diseases 18:1020– 1022.

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Mitchell CM, Hutton S, Myers GSA, Brunham R, Timms P. 2010. Chlamydia pneumoniae is genetically diverse in animals and appears to have crossed the host barrier to humans on (at least) two occasions. PloS Pathog 6: e1000903. Mutschmann F. 1998. Detection of Chlamydia psittaci in amphibians using an immunofluorescence test (IFT). Berliner und Münchener tierärztliche Wochenschrift 111: 187–189. Newcorner CE, Anver MR, Simmons JL, Wilcke RW, Nace GW. 1982. Spontaneous and experimental infections of Xenopus laevis with Chlamydia psittaci. Laboratory Animal Science 332: 680–686. Reed KD, Ruth GR, Meyer JA, Shukla SK. 2000. Chlamydia pneumonia infection in a breeding colony of African clawed frogs (Xenopus tropicalis). Emerging Infectious Disease 6: 196–199. Sharifian-Fard M, Pasmans F, Adriaensen C, Devisscher S, Adriaens T, Louette G, Martel A. 2011. Invasive bullfrogs constitute a reservoir of ranavirosis. Emerging Infectious Disease 17: 2371–2372. Vandewoestyne M, Van Nieuwerbrugh F, Van Hoofstat D, Deforce D. 2012. Evaluation of three DNA extraction protocols for forencsic STR typing after laser capture microdissection. Forensic Science International: Genetics 6: 258–262. Wright KM. 1996. Chlamydial infections of amphibians. Association of Reptilian and Amphibian Veterinarians 6: 8–9.

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Chironomidae bloodworms larvae as aquatic amphibian food

Mojdeh Sharifian Fard,1 Frank Pasmans,1 Connie Adriaensen,1 Gijs Du Laing,2 Geert Paul Jules Janssens,3 and An Martel1

1. Department of Pathology, Bacteriology and Avian Diseases, Ghent University Salisburylaan 133, B-9820 Merelbeke, Belgium 2. Laboratory of Analytical Chemistry and Applied Ecochemistry, Ghent University, Coupure Links 653, B-9000 Gent, Belgium 3. Department of Nutrition, Genetics and Ethology, Faculty of Veterinary Medicine, Ghent University, Heidestraat 19, B-9820 Merelbeke, Belgium

Adapted from Zoo Biology, 2014, 33(3): 221–227

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Abstract Different species of chironomids larvae (Diptera: Chironomidae) so-called bloodworms are widely distributed in the sediments of all types of freshwater habitats and considered as an important food source for amphibians. In our study, three species of Chironomidae (Baeotendipes noctivagus, Benthalia dissidens and Chironomus riparius) were identified in 23 samples of larvae from Belgium, Poland, Russia and Ukraine provided by a distributor in Belgium. We evaluated the suitability of these samples as amphibian food based on four different aspects: the likelihood of amphibian pathogens spreading, risk of heavy metal accumulation in amphibians, nutritive value, and risk of spreading of zoonotic bacteria (Salmonella, Campylobacter, and broad-spectrum beta-lactamase producing Enterobacteriaceae). We found neither zoonotic bacteria nor the amphibian pathogens ranaviruses and Batrachochytrium dendrobatidis in these samples. Our data showed that among the five heavy metals tested (Hg, Cu, Cd, Pb and Zn), the excess level of Pb in 2 samples and low content of Zn in 4 samples implicated potential risk of Pb accumulation and Zn inadequacy. Proximate nutritional analysis revealed that, although the high protein content in these samples could fulfill the protein needs, the wide range in lipid: protein ratio warrants further study, since this ratio will affect the amount and pathway of energy supply to the amphibians. Our study indicated that although environmentally-collected chironomids larvae may not be vectors of specific pathogens, they can be associated with nutritional imbalances and may also result in Pb bioaccumulation and Zn inadequacy in amphibians. Chironomidae larvae may thus not be recommended as single diet item for amphibians.

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1. Introduction Chironomidae are a family of nematoceran flies with a worldwide distribution. This large taxon of insects resembles mosquitoes but they lack the wing scales and elongate mouthparts of the Culicidae. In Europe, bloodworm is the common name for the aquatic larval stage of Chironomidae. However, this term is also attributed to Glyceridae (a family of polychaete worms) in some parts of the world (Kozloff, 1987). Chironomid larvae can be found in almost any aquatic or semiaquatic habitat and are important food items for both vertebrates and invertebrates (Armitage et al., 1995; Bat et al., 2001). Captive propagation of amphibians has gained attention in recent years, ex situ conservation being considered a last resort to safeguard many species from extinction (Stuart et al., 2004). Commercially available bloodworms are used in many aquatic amphibian species as important component of the captive diet. However, little is known about the nutrient composition of different species of chironomids larvae. In addition, feeding bloodworms may compromise the health of both the amphibians and their keepers. Indeed, since bloodworms feed on algae and detritus, they represent a possible reservoir of zoonotic agents (Salmonella, Vibrio cholerae, Campylobacter jejuni and Escherichia coli) (Rouf et al., 1993; Broza and Halpern, 2001; Moore et al., 2003) and of acquired antimicrobial resistance. Captive amphibian health may also be influenced by feeding bloodworms to amphibians, through possible transmission of ranavirus virions and Batrachochytrium dendrobatidis zoospores (causal agent of chytridiomycosis) (Gleason et al., 2008; Gray et al., 2009). Apart from infectious agents, bloodworms may contain high levels of heavy metals. Due to pollution and anthropogenic processes the prevalence of heavy metals such as lead (Pb), zinc (Zn), cadmium (Cd), mercury (Hg) and copper (Cu) has increased in aquatic environments (Blaustein et al., 2003), resulting in possible accumulation in bloodworms (Sharley et al., 2004). Amphibians are considered highly susceptible to heavy metal intoxications (Lefcort et al., 1998). In this study, we determined the identity and the nutritional value of a selection of bloodworms species originating from different countries, and the possible impact of the use of these bloodworms on human and amphibian health. Subsequently, their content of heavy metals and the presence of the following zoonotic and amphibian pathogens was determined: Salmonella spp., broad-spectrum beta-lactamase producing Enterobacteriaceae, Campylobacter spp., ranaviruses and Batrachochytrium dendrobatidis.

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2. Materials and Methods 2.1. Bloodworms Between October 2011 and April 2012, we collected chironomid larvae which were bought from a distributor in Belgium. This resulted in 23 bloodworm samples collected in Russia (18 samples), Poland (2 samples), Ukraine (1 sample) and Belgium (2 samples).

2.2. DNA preparation DNA was prepared for chironomid species identification and pathogen detection. We preserved all samples in 70% ethanol prior to DNA extraction. For the DNA preparation, the larvae were crushed to a fine powder in liquid nitrogen in a 1.5 mL Eppendorf tube and genomic DNA was extracted using the Bioline Isolate Genomic DNA Mini Kit (BIOLINE, London, UK). We stored extraction products at -20˚C.

2.3. PCR amplification to identify the chironomid species A 710-bp fragment of the mitochondrial cytochrome oxidase subunit (COI) was amplified as described by Sharley et al., (2004) using primers 911 (5΄TTTCTACAAATCATAAAGATATTGG3΄) as forward primer and HCO2198 (5΄TAAACTTCAGGGTGACCAAAAAATCA3΄) as reverse primer. The PCR products were purified by PCR purification kit (QIAGEN, GmbH, Hilden, Germany) according to manufacturer’s instructions and se uenced with the same primers used in the PCR on an automated DNA sequencer (ABI3700, Applied Biosystems). The sequence data were assembled and aligned using LICOR AlignIR Version 2. The obtained data were then compared to those published in GenBank through BLAST network service (http://www.ncbi.nlm.nih.gov/BLAST).

2.4. Isolation of Salmonella, Campylobacter and broad-spectrum beta-lactamase producing Enterobacteriaceae The fresh larvae were homogenized thoroughly by grinding and then we divided into three subsamples for detection of Salmonella spp., broad-spectrum beta-lactamase producing Enterobacteriaceae and Campylobacter spp. We used sterile procedures and instruments for each sample throughout the study to prevent the cross-contamination of the samples. We investigated the presence of Salmonella spp. according to the EN ISO 6579:2002 standard method which was modified according to the recommendations of the Community

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Reference Laboratory for Salmonella in Bilthoven, the Netherlands. Briefly, pre-enrichment was done by incubation of the samples in buffered peptone water (Oxoid, Basingstoke, UK) during 18 h at 37°C. From the pre-enrichment solution, 3 droplets were inoculated in a Rappaport-Vassiliadis broth (Difco, Becton, Dickinson and Co., New York, US) and Tetrathionate broth (Oxoid). Then they were incubated for 24 h at 37°C. Samples from both broths were plated on Brilliant Green agar (Oxoid) and Xylose-Lysine-Deoxycholate agar (Oxoid), followed by incubation for 24 h at 37 °C. For the isolation of broad-spectrum beta-lactamase producing Enterobacteriaceae, each homogenized sample was directly plated onto McConkey agar (Oxoid) containing 8 µg/ml ceftiofur and was incubated for 24 h at 37°C. For detection of Campylobacter spp. 1 gram of each sample was diluted in nutrient broth No.2 (Oxoid) supplemented with Modified Preston Campylobacter-selective supplement (Oxoid) and Campylobacter-specific growth supplement (Oxoid) and incubated for 24 h at 42°C under microaerobic conditions. Then diluted samples were plated onto modified Charcoal Cefoperazone Deoxycholate agar (Oxoid) plates, supplemented with CCDA selective supplement (Oxoid) and Campylobacter-specific growth supplement (Oxoid). The plates were incubated for 48 h at 42°C under microaerobic conditions.

2.5. Detection of zoonotic and amphibian pathogens using PCR In parallel to bacteriological cultures, we tested all chironomid larvae samples for the presence of Salmonella, Campylobacter, ranaviruses, and B. dendrobatidis by PCR. A PCR to detect the presence of Salmonella was performed as described by Rahn et al., (1992) using the invA gene primers 5´GTGAAATTATCGCCACGTTCGGGCAA3´ and 5´TCATCGCACCGTCAAAGGAACC3´. A negative control (HPLC water) and positive control (DNA extracted from Salmonella Typhimurium DAB69) were used in each PCR run. For the detection of Campylobacter a PCR was performed as described by Wegmuller et al., (1993) using Campylobacter jejuni flaA and flaB gene primers 5´GCTCAAAGTGGTTCTTATGCNATGG3´ and 5´GCTGCGGAGTTCATTC TAAGACC3´. A negative control (HPLC water) and a positive control (DNA extracted from C. jejuni KC40) were included in each PCR run. For the detection of ranaviruses, a PCR reaction was performed following the protocol of Mao et al., (1997) using the major capsid protein (MCP) of FV3 gene primers 5´GACTTGGCCACTTATGAC3´ and 5´GTCTCTGGAGAAGAAGAA3´. Negative control (HPLC water) and positive control (DNA extracted from an infected and confirmed ranavirus-

125 STUDY 4 positive tadpole) served as controls for the PCR runs. Amplification products were run on 1.5% agarose gel, immersed in TAE buffer (40 mM Tris–Acetat, 1 mM EDTA) stained with Ethidium Bromide 0.5 μg/mL and visualized with UV transiluminator. The presence of B. dendrobatidis was investigated with quantitative PCR (qPCR) described by Boyle et al., (2004). Subsequently, DNA samples were diluted 1:10 and qPCR assays were performed in duplicate on a CFX96 Real Time System (BioRad Laboratories, Hercules, CA, USA). We used ITS1-3 Chytr (5-CCTTGATATAATACAGTGTGCCATATGTC-3) for the forward primer, 5.8S Chytr (5-AGCCAAGAGATCCGTTGTCAA-3) for the reverse primer and Chytr MGB2 for the TaqMan Probe (5-6FAM CGAGTCGAACAAAAT MGBNFQ-3). One positive control sample containing B. dendrobatidis DNA (JEL423) and three negative control samples with HPLC water were included for each assay. To control and estimate inhibition, a subset of samples negative for the presence of B. dendrobatidis (n = 20) was retested under the same conditions as described above, but with an exogenous internal positive control ( IC™ probe, Life technologies, Austin, TX, USA) included as described by Hyatt et al., (2007).

2.6. Determination of heavy metal content We analyzed concentrations of lead (Pb), zinc (Zn), cadmium (Cd), mercury (Hg) and copper (Cu) using the method described by Tack et al., (2000). For this purpose, 7 ml ultrapure 65% HNO3 was added to 1 gram of the larvae in a 100 ml Pyrex beaker covered with a watch-glass. The suspension was heated up to 150ºC for 2 hours. Then it was treated with 4 ml 30% H2O2. After cooling, the solution was transferred to a 50 ml flask and diluted to the mark with distilled de-ionized water. Determination of Cd, Cu, Zn and Pb in the solution was performed by inductively coupled plasma optical emission spectrometry (ICP- OES, Varian Vista MPX, Palo Alto, CA, USA) when results were below detection limit of ICP-OES. Mercury was measured using a cold vapor atomic absorption mercury analyzer (CV-AAS, CETAC QuickTrace M-7500, Omaha, NE, USA).

2.7. Nutritional analysis All chironomid larvae samples were stored frozen at -20˚C prior proximate analysis. To determine nutritional analysis, each sample was individually analyzed. We measured dry matter (DM) as performed by De La Noue and Choubert (1985). Larvae were dried at 95˚C for 24 hours until a constant weight was achieved. Ash contents were determined as described

126 STUDY 4 by Bogut et al., (2007) burning the larvae at 550˚C for 4 hours in a muffle furnace. We determined crude protein (CP) content according to Kjeldahl × 6.25 methods using the Kjel- Foss, and crude fat (CF) content by Soxhlet methods. Neutral detergent fibre (NDF) and acid detergent fibre (ADF) were determined according to Association of Official Analytical Chemist (AOAC, 1995) methods.

2.8. Statistical analysis We evaluated the overall difference between species for each variable by using either an ANOVA or the non-parametric Kruskal-Wallis test. For each variable, an ANOVA model was built. Normality of the residuals for each model was evaluated using a QQ plot. Models without normally distributed residuals were disregarded, and a Kruskal-Wallis test was used instead for those variables. When applicable, a Dunnett T3 pairwise comparison test was carried out as well. This post-hoc test could deal with unequal variances and sample sizes, while being more conservative than e.g. the Games-Howell post-hoc test. It must be noted that due to the unequal sample sizes, the Kruskal Wallis test might be overly optimistic (Zimmerman, 2000).

3. RESULTS 3.1. Identification of the larvae Fourteen samples originating from Russia and the only sample from Ukraine consisted of Baeotendipes noctivagus. The samples from Belgium (two samples), Poland (two samples) and one sample from Russia contained the species Chironomus riparius. Three samples from Russia belonged to the species Benthalia dissidens.

3.2. Zoonotic and amphibian pathogens Cultures for Salmonella, Campylobacter and broad-spectrum beta-lactamase producing Enterobacteriaceae were all negative. The negative results for Salmonella and Campylobacter were confirmed by PCR. The PCR’s for the amphibian pathogens B. dendrobatidis and ranaviruses were all negative.

3.3. Heavy metal content For the three species identified, B. noctivagus (n=15), C. riparius (n=5) and B. dissidens (n= 3), the concentrations of heavy metals varied between 0.0007 and 0.0790 µg/g for Hg, 2- 17 µg/g for Cu, <0.003-0.351 µg/g for Cd, 0.6-9.9 µg/g for Pb and 9-59 µg/g for Zn on a dry

127 STUDY 4 matter basis (DMB). The residual analysis of the ANOVA models showed for all metals a moderate to strong deviation from normality. Therefore, we evaluated our data using Kruskal-Wallis. The Hg concentrations differed significantly between groups (p=0.007). The boxplot (Figure 1) points towards lower concentrations for C. riparius compared to B. noctivagus. In addition, C. riparius contained higher concentrations of Zn compared to the two other species (p=0.007). The distribution of the measurements within each species is given in Figure 1.

Figure 1. Box- and point plots for trace metals (Zn and Cu) and xenobiotic heavy metals (Hg, Cd and Pb) concentrations (µg/g) per chironomid species on a dry matter basis (DMB). The point line indicates the maximum Pb levels for fish (as typical aquatic animals) food (5 µg/g) and exhibits two samples are above the maximum limits. The black line indicates the minimum Zn requirement as trace metal (15 µg/g) in fish diet and shows four samples are below the limit. Other metal values were in acceptable limits in fish food.

3.4. Nutritive value The QQ plots showed in general an acceptable normality for the residual values. Hence, we could use the ANOVA models for comparison of the different species. Differences between species were found for CP (p<0.001), ADF (p=0.007), CF (p<0.001) and Ash (p=0.001). According to the post hoc tests, C. riparius differed from B. noctiavagus and B. dissidens by a significantly higher content of both crude protein and crude fat (p<0.05) and a significantly lower content of ADF (p< 0.05). B. dissidens contained significantly less crude fat than the other two species (p<0.05). However, due to the low numbers of B. dissidens, it should be careful drawing firm conclusions from these results. The distribution of the nutrient values per chironomid species are illustrated in Figure 2.

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Figure 2. Box- and point plots for nutrients measures per chironomid species as percent on a dry matter basis (DMB) within species. Black lines indicate the average value of each nutrient including 19.95±2.06, 52.11±4.73, 16.96±2.65, 19.58±6.19, 4.50±2.68, and 25.95±3.78 for DM, CP, ADF, Ash, CF, and NDF, respectively. DM: Dry Matter, CP: Crud Protein, ADF: Acid Detergent Fiber, CF: Crud Fat, NDF: Neutral Detergent Fiber.

4. Discussion In our samples, commercially available “bloodworms” comprise at least three different species of Chironomidae larvae (B. noctivagus, C. riparius and B. dissidens) with B. noctivagus as the predominant species. Since spatial patterns of species diversity can change over spatial scales, the pattern observed in this study might be different from those found in other parts of the world. In contrast with previous studies (Rouf et al., 1993; Broza and Halpern, 2001; Moore et al., 2003), none of the tested samples were positive for Salmonella, Campylobacter, broad- spectrum beta-lactamase producing Enterobacteriaceae, ranaviruses or B. dendrobatidis.

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Thus, the use of bloodworms appears to pose a minor risk regarding spread of these pathogens to humans or amphibians. However, samples used in this study only represent a snapshot of possible contamination, not taking into account for example seasonal fluctuations in disease dynamics (Bosch et al., 2007; Todd-Thompson, 2010). We compared the heavy metal concentrations in the bloodworms with the maximum metal content of Hg (0.1µg/g) (The EFSA Journal, 2008), Cu (35µg/g) (European commission Directive 70/524/EEC, 2003), Cd (2µg/g) (EU in Directive 2005/87/EC), Pb (5µg/g) (EU in Directive 2005/87/EC) and Zn (250µg/g) (European commission Directive 70/524/EEC, 2003) in fish (as typical aquatic animals) diet. Our results revealed that the concentration of Pb in two samples exceeded the maximum metal levels. However, four samples contained a lower concentration of Zn than the minimum requirement level (15µg/g) in fish diet (Watanabe et al., 1997). The excessive levels of Pb and low contents of Zn render the sole feeding of commercial bloodworms to captive amphibians questionable. Hence, feeding a variety of prey items is advisable to overcome Zn inadequacy and to maintain balanced physiological metals content for amphibians. Since dietary zinc has an antagonistic effect on copper absorption (Evans et al., 1970) leading to decreased copper status in organisms, the high concentrations of Zn in C. riparius suggests a decreased bio-availability of Cu in this species. The different load of heavy metals in chironomids larvae most probably correlates with content of metals in water and sediments. The synergistic effect of different physical and chemical factors such as temperature and hardness of water, sediment pH, exposure time, chemical form and availability of metals might promote accumulation of heavy metals into chironomids larvae (Bhattacharya et al., 2006; Lagrana et al., 2010). To eliminate or reduce the environmental impacts (e.g. heavy metals load), captive rearing of bloodworms in controlled environments is advisable, though there are great difficulties and laborious in keeping cultures going and failures can be occurred betimes (Galtsoff et al., 1937; Richardson et al., 2005). Although amphibians can detoxify both physiological (such as zinc and copper) and xenobiotic (such as cadmium, mercury and lead) heavy metals through metallothionein proteins, excess quantities of metals can induce toxicity (acute or chronic) in amphibian species (Muller et al., 1993; Saint-Jacques and Séguin, 1993; Lance et al., 2011). Pb intoxication can lead to malformations, delayed developmental rate and decreased muscle tone (Horne and Dunson, 1995). When comparing our results with those from previously published work (Sugden, 1973;

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Armitage, 1995; Habib et al., 1997; Habashy, 2005; Bogut et al., 2007; Thipkonglars et al., 2010; Rajabipour et al., 2011; Pasmans et al., 2012) the protein content of chironomid larvae varies markedly, between 31 and 67% on a dry matter basis. Similarly, fat and ADF content showed considerable variation between the different studies (Armitage, 1995; Bernard et al., 1997; Habib et al., 1997; Habashy, 2005; Bogut et al., 2007; Thipkonglars et al., 2010; Pasmans et al., 2012), from 3 to 14% and 4 to 17% on a dry matter basis, respectively. The nitrogen in chitin (exoskeleton) estimated by ADF is likely not contributing to the dietary protein supply of insectivore amphibians, hence the ADF content indicates how much of the analysed crude protein is actually non-protein nitrogen. It is therefore likely that C. riparius provides more dietary protein due to the lower percentage of ADF in comparison with the two other species (P<0.05). Differences within or between species or studies may be caused by intrinsic differences in composition and/or by the environmental context. Indeed, the proximate composition of larvae samples may depend on the larvae age, composition of earthpond water, food availability and food quality and quantity (Mackey, 1977; Vos et al., 2000; Habashy, 2005; Bogut et al., 2007; Rajabipour et al., 2011). In addition, environmental factors such as temperature, photoperiod, pH, oxygen content and biotic interaction can affect growth and consequently nutritional composition in bloodworms (Maier et al., 1990; Tokeshi, 1995). Since a suitable diet for insectivore amphibians contains 30% to 60% proteins (McWilliams, 2008), our analysis shows chironomid with the average protein value 52.1%±4.73 to be high in protein content. Whereas the dietary fat fractions for amphibians naturally range from less than 10% to more than 30% (McWilliams, 2008) the obtained low fat contents in this (4.5%) and other studies (between 3 and 14%) indicates that feeding environmentally-collected chironomids larvae might not comply needs for amphibian diet program. Since lipid content largely determines the energy content, hence low lipid content might not ensure the needed dietary energy density. However, McWilliams (2008) provided basic information on what to feed captive amphibians, this area of study is still lagging behind the disciplines of mammalian and avian nutrition and further studies should fortify knowledge of amphibian nutrition involving different species and different developmental growth stages.

5. Conclusion In our study we found that although chironomids larvae can meet high protein needs in amphibian diet program with apparently low risk of transmission of infectious organisms, relatively high levels of Pb and low levels of Zn combined with low fat content renders

131 STUDY 4 feeding environmentally-collected chironomids larvae as single diet item to amphibians unsuitable.

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6. Refrences Armitage PD. 1995. Chironomidae as food. In: Armitage PD, Cranston PS, Pinder LCV, editors. The Chironomidae: Biology and Ecology of Non-Biting Midges. Chapman and Hall, London.UK. pp 423-435. AOAC (Association of Official Analytical Chemists). 1990. Official Methods of Analysis 15th edn. Association of Official Analytical Chemists. Arlington, AV, 1298 pp. Bat L, Akbulut M. 2001. Studies on sediment toxicity bioassays using Chironomus thummi K., 1911 Larvae. Turkish Journal of Zoology 25:87-93. Bernard JB, Allen ME, Ullrey DE. 1997. Feeding Captive Insectivorous Animals: Nutrition Aspects of Insects as Food. Nutrition Advisory Group Handbook Fact Sheet 005. Bhattacharya G, Sadhu AK, Mazumbar A, Majumdar U, Chaudhuri PK, Ali A. 2006. Assessment of impact of heavy metals on the communities and morphological deformities of Chironomidae larvae in the river Damodar (India, West Bengal). Supplementa ad Acta Hydrobiologica 8:21-32. Blaustein AR, Romansic JM, Kiesecker JM, Hatch AC. 2003. Ultraviolet radiation, toxic chemicals and amphibian population declines. Diversity and Distributions 9:23-140. Bogut I, Has-Schon E, Adamek Z, Rajković , Galović D. 2007. Chironomus plumosus larvae - a suitable nutrient for freshwater farmed fish. Poljoprivreda 13(1):159-162. Bosch J, Carrascal LM, Duran L, Walker S, Fisher MC. 2007. Climate change and chytridiomycosis in a montane area of Central Spain: Is there a link? Proceedings of the Royal Society of London, Series B 274:253-260 Boyle DB, Olsen V, Morgan JAT, Hyatt AD. 2004. Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman PCR assay. Diseases of Aquatic Organisms 68:47–50. Broza M, Halpern M. 2001. Chironomid egg masses and Vibrio cholerae. Nature 412:40. De La Noue J, Choubert G. 1985. Apparent digestibility of invertebrate biomass by rainbow trout. Aquaculture 50:103-112. EFSA (The European Food Safety Authority Journal). 2008. Opinion of the Scientific Panel on Contaminants in the Food chain on a request from the European Commission on mercury as undesirable substance in feed 654:1-74. EU (European Union) Directive 2005/87/EC. Amending Annex I to Directive 2002/32/EC of the European Parliament and of the Council on undesirable substances in animal feed as regards lead, fluorine and cadmium. European commission Directive 70/524/EEC. 2003. Committee for Animal Nutrition on the

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use of copper in feeding stuffs. European commission Directive 70/524/EEC. 2003. Opinion of the Scientific Committee for Animal Nutrition on the use of zinc in feeding stuffs. Evans GW, Majors PF, Comatzer WE. 1970. Mechanism of cadmium and zinc antagonism of copper metaboUsm. Biochemical and Biophysical Research Communications 40:142- 8. Fort DJ, Propst TL, Stover EL, Helgen JC, Levey R, Gallagher K, Burkhart JG. 1999. Effects of pond water, sediment, and sediment extracts from Minnesota and Vermont on early development and metamorphosis in Xenopus. Environmental Toxicology and Pharmacology 18:2305–15. Galtsoff PS, Welch PL, Lutz FE. 1937. Culture Methods for Invertebrate Animals. Dover Publications Inc. 590 pp. Gleason HF, Kagami M, Lefevre E, Sime-Ngando T. 2008. The ecology of chytrids in aquatic ecosystems: roles in food web dynamics. Fungal Biology Reviews Journal 22:17-25. Gray MJ, Debra DL, Hoverman JT. 2009. Ecology and pathology of amphibian ranaviruses. Diseases of Aquatic Organisms 87:243–266. Habashy MM. 2005. Culture of chironomid larvae (Insecta- Diptera- Chironomidae) under different feeding systems. Egyptian Journal of Aquatic Research 31: 403-418. Habib MAB, Yusoff FM, Phang SM, Ang KJ, Mohamed S. 1997. Nutritional values of chironomid larvae grown in palm oil mill effluent and algal culture. Aquaculture 158:95- 105. Hald B, Skovgård H, Pedersen K, Bunkenborg H. 2008. Influxed insects as vectors for Campylobacter jejuni and Campylobacter coli in Danish broiler houses. Poultry Science 87:1428-1434. Horne MT, Dunson WA. 1995. Effects of low pH, metals, and water hardness on larval amphibians. Archives of Environmental Contamination and Toxicology 29:500-505. Kozloff EN. 1987. Marine Invertebrates of the Pacific Northwest. University of Washington Press, Seattle, WA. 511 pp. ISBN 0-295-96530-4. Lagrana C, Apodaca D, David CP. 2010. Response of chironomids to Cu2+, Cd2+ and Pb2+ metal ions using the 96-hour toxicity test and evaluating different factors that affect metal accumulation. Journal of Trends in Chemistry 1(1):8-13. Lance SL, Erickson MR, Flynn RW, Mills GL, Tuberville TD, Scott DE. 2012. Effects of chronic copper exposure on development and survival in the southern leopard frog (Lithobates [Rana] sphenocephalus). Environmental Toxicology and Chemistry 31(7): 1587-1594.

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Lefcort H, Meguire RA, Wilson LH, Ettinger WF. 1998. Heavy metals alter the survival, growth, metamorphosis, and antipredatory behavior of Columbia spotted frog (Rana luteiventris) tadpoles. Environmental Contamination and Toxicology 35(3):447-56. Mackey AP. 1977. Quantitative studies on the Chironomidae (Diptera) of the River Thames and Kennet. 111. The Nuphar Zone. Archive Hydrobiologia 79:62-102. Maier KJ, Kosalwat P, Knight AW. 1990. Culture of Chironomus decorus (Diptera: Chironomidae) and the effect of temperature on its life history. Environmental Entomology 19: 1681-8. Mao J, Hedrick RP, Chinchar VG. 1997. Molecular characterization, sequence analysis, and taxonomic position of newly isolated fish iridoviruses. Virology 229(1):212-20. McWilliams DA. 2008. Nutrition recommendations for some captive amphibian species (Anura and Caudata). The Canadian Association of Zoos and Aquariums Nutrition Advisory and Research Group (GARN-NARG). Moore BC, Martinez E, Gay JM, Rice DH. 2003. Survival of Salmonella enterica in freshwater and sediments and transmission by the aquatic midge Chironomus tentans (Chironomidae: Diptera). Applied and Environmental Microbiology 69:4556-60. Muller JP, Wouters-Tyrou D, Erraiss NE, Vedel M, Touzet N, Mesnard J, Sautière P, Wegnez M. 1993. Molecular cloning and expression of a metallothionein mRNA in Xenopus laevis. DNA Cell Biology 12:341–349. Pasmans F, Janssens GPJ, Sparreboom M, Jiang J, Nishikawa K. 2012. Reproduction, development, and growth response to captive diets in the shangcheng stout salamander, Pachyhynobius shangchengensis (Amphibia, Urodela, Hynobiidae). Asian Herpetological Research 3(3):192–197. Rahn K, De Grandis SA, Clarke RC, McEwen SA, Galán JE, Ginocchio C, Curtiss R .3rd, Gyles CL. 1992. Amplification of an invA gene sequence of Salmonella typhimurium by polymerase chain reaction as a specific method of detection of Salmonella. Molecular and Cellular Probes 6(4):271-9. Rajabipour F, Mashaii N, Saresangi H, Bitaraf A, Mohammadi M, Sahragard A. 2011. Chironomus Aprilinus Meigen, 1830, production in underground brackish waters of Iran. Academic Journal of Entomology 4(2):41-46. Richardson BM, Stence CP, Baldwin MW. 2005. Development of a captive brood stock program for Atlantic sturgeon (Acipenser oxyrhinchus) in Maryland. Progress Report for U.S. Fish & Wildlife Service State Wildlife. Pp 43. Rouf MA, Rigney MM. 1993. Bacterial florae in larvae of the lake fly Chironomus plumosus.

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Applied and Environmental Microbiology 59(4):1236–41. Saint-Jacques E, Séguin C. 1993. Cloning and nucleotide sequence of a complementary DNA encoding Xenopus laevis metallothionein: mRNA accumulation in response to heavy metals. DNA Cell Biology 12:329–40. Sharley DJ, Pettigrove V, Parson YM. 2004. Molecular identification of Chironomus spp. (Diptera) for biomonitoring of aquatic ecosystems. Australian Journal of Entomology 43: 359-65. Stuart SN, Chanson JS, Cox NA, Young BE, Rodrigues ASL, Fischman DL, Waller RW. 2004. Status and trends of amphibian declines and extinctions worldwide. Science 306 (5702): 1783-6. Sugden LG. 1973. Feeding ecology of pintail, gadwall, American widgeon and lesser scaup ducklings in southern Alberta. Canadian Wildlife Services Report 24:45. Tack FMG, Bogaert N, Verloo MG, Hendrickx F, Maelfait JP, Mertens J. 2000. Determination of Cd, Cu, Pb and Zn in woodlouse (Oniscus asellus). International Journal of Environmental Analytical Chemistry 78:149-58. Thipkonglars N, Taparhudee W, Kaewnern M, Lawonyawut K. 2010. Cold preservation of chironomid larvae (Chironomus fuscipes Yamamoto, 1990): nutritional value and potential for climbing perch (Anabas testudineaus Bloch, 1792) larval nursing. Kasetsart University Fisheries Research Bulletin 34(2):1-13. Todd-Thompson M. 2010. Seasonality, variation in species prevalence, and localized disease for Ranavirus in Cades cove (Great Smoky Mountains National Park) amphibians. University of Tennessee; Knoxville, TN, USA: Master Thesis, Available online: http://trace.tennessee.edu/utk_gradthes/665 (accessed on 17 November 2011). Tokeshi M. 1995. Life cycles and population dynamics In: Armitage PD, Cranston PS, Pinder LCV, editors. The Chironomidae: Biology and Ecology of Non biting Midges. Chapman and Hall. London UK. Pp225-250. Vos JH, Ooijevaar MAG, Postma JE, Admiraal W. 2000. Interaction between food availability and food quality during growth of early instar chironomid larvae. Journal of North American Benthological Society 19(1):158-68. Watanabe T, Kiron V, Satoh S. 1997. Trace minerals in fish nutrition. Aquaculture 151:185- 207. Wegmuller B, Luthy J, Candrian U. 1993. Direct polymerase chain reaction of Campylobacter jejuni and Campylobacter coli in raw milk and dairy products. Applied and Environmental Microbiology 59:2161–5.

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Wilson IG. 1997. Inhibition and facilitation of nucleic acid amplification. Applied and Environmental Microbiology 63:3741-51. Zimmerman DW. 2000. Statistical significance levels of nonparametric tests biased by heterogeneous variances of treatment groups, The Journal of General Psychology 127(4): 354-64.

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Migrating common toads (Bufo bufo) in rural temperate regions: reservoirs of Salmonella?

Mojdeh Sharifian Fard, Frank Pasmans and An Martel

Department of Pathology, Bacteriology and Avian Diseases, Ghent University, Salisburylaan 133, B-9820 Merelbeke, Belgium

Adapted from Journal of Wildlife Diseases, 2014, 50(2): 326-9

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Abstract Salmonella infections in amphibians are supposedly highly prevalent. Migrating common amphibian species in cultivated areas such as common toads (Bufo bufo) may thus promote spread and zoonotic transfer of Salmonella to humans both indirectly by crop and livestock contamination and by direct contact. Between February and April 2011, the intestinal content of 1,740 samples of road killed migrating common toads in five Flemish provinces of Belgium was examined for the presence of Salmonella using bacterial culture and PCR. All the samples were negative. These results suggest that the role of migrating common toads in maintaining the infection cycle of Salmonella in northern European temperate regions is negligible.

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Salmonella infections are amongst the most important bacterial zoonoses with an incidence of 23.7 confirmed cases per 100,000 human populations in the European Union (EFSA and ECDC, 2011). Infected livestock and crops are prime sources of Salmonella infection in man. Animals (e.g. rodents, birds, and insects) are considered important vectors of Salmonella, maintaining infection cycles in agricultural areas (Wang et al., 2011; Umali et al., 2012). Although amphibians are generally considered to exert a positive effect in agriculture by consuming large numbers of invertebrates (Stewarta and Woolbright, 1996), they are also considered asymptomatic reservoirs for Salmonella serotypes such as Enteritidis, Rubislaw, Saintpaul, Montevideo and Newport (O’Shea et al., 1990; Chambers and Hulse, 2006; Drake et al., 2012). Since common toads (Bufo bufo) are among the most abundant amphibians in rural areas and on farms in northern Europe, and exhibit marked migratory activity during the reproductive season, they could be important vectors of Salmonella. Besides indirect transfer of Salmonella through contamination of crop and livestock, several outbreaks of human salmonellosis due to S. enterica serotypes Javiana and Typhimurium, have been directly associated with amphibian contact (Srikantiah et al., 2001; Clarkson et al., 2010). In many northern European countries, animal conservation groups (often involving children) participate in amphibian safeguarding activities during the spring migration. Thus, infected toads may pose the risk of direct Salmonella transfer to humans as well. Since Salmonella prevalence studies have been mostly performed in regions with warm climates (O’Shea et al., 1990; Drake et al., 2012), very little is known regarding the role of amphibians as carriers of Salmonella in temperate regions. Indeed Pasmans et al. (2013) hypothesized that the significance of ectothermic vertebrates as Salmonella reservoirs is limited to those species that provide sufficiently high body temperatures for successful persistent host colonization. Thus, our hypothesis is that amphibians from temperate climates with low thermal preferences are no significant reservoir hosts for Salmonella. The aim of this study was, therefore, to investigate the prevalence of Salmonella carriers among migrating common toads (Bufo bufo). Between 27 February and 7 April 2011, 1,740 road-killed common toads (Bufo bufo) were collected from 104 migration sites in all five Flemish provinces including Antwerp, East Flanders, Flemish Brabant, Limburg and West-Flanders (Figure 1).

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Figure 1. Sampling localities of common toads (Bufo bufo) in Flemish regions of Belgium

Specified meteorological data in the time period of this study were provided by the Royal Meteorological Institute of Belgium (Figure 3). Each sample was put in a sterile plastic bag and sterile procedures and instruments were used throughout the study to prevent the cross-contamination of the samples. Toads were dissected, and the intestinal content from the small and large intestine of each individual was thoroughly homogenized by grinding. The homogenate was placed in 2 mL microcentrifuge tubes and stored at -20°C. The presence of Salmonella was investigated according to the EN ISO 6579:2002 standard method, which was modified according to the recommendations of the Community Reference Laboratory for Salmonella in Bilthoven, the Netherlands. Briefly, pre-enrichment was done by incubation of the samples in buffered peptone water (Oxoid, Basingstoke, UK) during 18 h at 37°C. From the pre-enrichment solution, 3 droplets were inoculated in a Rappaport-Vassiliadis broth (Difco, Becton, Dickinson and Co., New York, US) and Tetrathionate broth (Oxoid). Then they were incubated for 24 h at 37°C. Samples from both broths were plated on Brilliant Green agar (Oxoid) and Xylose-Lysine-Deoxycholate agar (Oxoid), followed by incubation

144 STUDY 5 for 24 h at 37 °C. Genomic DNA was prepared using the QIAamp DNA Stool Mini Kit (QIAGEN, GmbH, Hilden, Germany). Extraction products were stored at -20 ˚C. A diagnostic PCR to identify Salmonella was performed as described by Chiu and Ou (1996) to amplify a 243 bp segment of the Salmonella invA gene using primer pair invA-1 (5´-ACAGTGCTCGTTTACGACCTGAAT) and invA-2 (5´-AGACGACTGGTACTGATCGATAAT). The negative control was a PCR mixture without DNA template, while the positive control was with DNA extracted from Salmonella Typhimurium DAB69. To determine the detection limit for the PCR assay, a dilution experiment was performed in the laboratory using the modified protocol proposed by Pathmanathan et al., (2003). Salmonella serotype Typhimurium strain DAB69 was grown in Brain Heart Infusion (BHI) broth at 37 ˚C overnight, and the Salmonella concentration was determined by plating serial dilutions onto BHI agar and performing plate counts after 24 hr of growth at 37 ˚C. A quantity of 100 μl of each of the serial 10-fold dilutions of the original overnight culture was added to separate tubes containing 1 mg Salmonella-negative frog feces. The DNA was extracted from 100 μl of the spiked feces as described in an earlier section, and PCR was then performed on these extracts as described in the preceding text. The minimum detection limit of the invA PCR was between 30 and 300 cells. The results showed that all the 1,740 samples were negative for Salmonella. No Salmonella was isolated or detected using PCR. The reason for absence of Salmonella might be associated with thermal growth limits of Salmonella. As shown in Figure 2, the Salmonella growth rate substantially reduces at temperatures of 10 ºC. The mean environmental temperature during the collection period was below 10°C and varied between 7.6 and 8.3°C (Figure 3). Besides, toads are nocturnal, meaning that they won’t have been exposed to the maximum day temperatures. Since in this study only adult, migrating toads were investigated, we cannot rule out that other life stages are carriers for Salmonella. Because these life stages are infrequently handled by people, they do not provide a high risk.

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Figure 2. Predicted growth rates of Salmonella at different temperatures by the models of Oscar (2006) and Juneja et al., (2007) in Dominguez Risco (2009).

Figure 3. Environmental minimum, maximum, and mean temperatures for the five investigate provinces of Flanders between 27 February and 7 April 2011.

The low thermal preferences of European common toads (Reading, 2003) could preclude successful and persistent colonization of Salmonella. We thus suggest that amphibians with low thermal preferences do not provide a suitable niche for Salmonella, which renders their role in maintaining infection cycles negligible.

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Acknowledgment We thank Natuurpunt–Hyla and Natuurpunt volunteers who helped with sampling the road victims.

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References Chambers DL, Hulse AC. 2006. Salmonella serovars in the herpetofauna of Indiana County, Pennsylvania. Applied and Environmental Microbiology 72:3771–3. Chiu CH, Ou JT. 1996. Rapid identification of Salmonella serovars in feces by specific detection of virulence genes, invA and spvC, by an enrichment broth culture culture- multiplex PCR combination assay. Journal of Clinical Microbiology 34:2619-22. Clarkson LS, Tobin D'angelo M, Shuler C, Hanna S, Benson J, Voetsch AC. 2010. Sporadic Salmonella enterica serotype Javiana infections in Georgia and Tennessee: a hypothesis-generating study. Epidemiology and Infection 138(3):340-6. Drake M, Amadi V, Zieger U, Johnson R, Hariharan H. 2012. Prevalence of Salmonella spp. in cane toads (Bufo marinus) from Grenada, West Indies, and their antimicrobial susceptibility. Zoonoses and Public Health. 60(6):437-41. Dominguez Risco SA. 2009. Risk factors influencing the growth and survival of salmonella on poultry products. PhD Dissertation. Rutgers, the State University of New Jersey. 126 pp. EFSA (European Food Safety Authority) and ECDC (European Centre for Disease Prevention and Control). 2011. The European Union summary report on trends and sources of zoonoses, zoonotic agents and food-borne outbreaks in 2009. Journal of European Food Safety Authority 9(3):2090. Juneja VK, Melendres MV, Huang LH, Gumudavelli V, Subbiah J, Thippareddi H. 2007. Modeling the effect of temperature on growth of Salmonella in chicken. Food Microbiology 24(4):328–35. O’shea P, Speare R, Thomas AD. 1990. Salmonella serotypes from the cane toad, Bufo marinus. Australian Veterinary Journal 67:310. Oscar TP. 2006. Validation of a tertiary model for predicting variation of Salmonella Typhimurium dt104 (ATCC 700408) growth from a low initial density on ground chicken breast meat with a competitive Microflora. Journal of food protection 69(9):2048-57. Pasmans F, Boyen F, Haesebrouck F. 2013. Salmonella in exotic pet animals. In: Barrow, P.A. (ed.) Salmonella in domestic animals. CABI publishing. Pp:337-50. Reading CJ. 2003. The effects of variation in climatic temperature (1980–2001) on breeding activity and tadpole stage duration in the common toad, Bufo bufo. Science of the Total Environment 310:231–6. Srikantiah P, Lay JC, Hand S, Crump JA, Campbell J, Van Duyne MS, Bishop R, Middendor

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R, Currier M, Mead PS, Mølbak K. 2004. Salmonella enterica serotype Javiana infections associated with amphibian contact, Mississippi, 2001. Epidemiology and Infection 132:273–81. Stewart MM, Woolbright LL. 1996. Amphibians. In: Reagan DP and Waide RP (Eds). The food web of a tropical rainforest. Chicago, IL: University of Chicago Press. Pathmanathan SG, Cardona-Castro N, Sánchez-Jiménez MM, Correa-Ochoa MM, Puthucheary SD, Thong KL. 2003. Simple and rapid detection of Salmonella strains by direct PCR amplification of the hilA gene. Journal of Medical Microbiology 52:773–6. Umali DV, Lapuz RR, Suzuki T, Shirota K, Katoh H. 2012. Transmission and shedding patterns of Salmonella in naturally infected captive wild roof rats (Rattus rattus) from a Salmonella-contaminated layer farm. Avian Diseases 56(2):288-94. Wang YC, Chang YC, Chuang HL, Chiu CC, Yeh KS, Chang CC, Hsuan SL, Lin WH, Chen TH. 2011. Transmission of Salmonella between swine farms by the housefly (Musca domestica). Journal of food protection 74(6):1012-6.

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The absence of zoonotic agents in invasive bullfrogs (Lithobates catesbeianus) in Belgium and The Netherlands

An Martel,1 Connie Adriaensen,1 Mojdeh Sharifian-Fard,1 Annemarieke Spitzen-van der Sluijs,2 Gerald Louette,3 Kristof Baert,3 Ben Crombaghs,4 Jeroen Dewulf,5 and Frank Pasmans1

1. Department of Pathology, Bacteriology and Avian Diseases, Ghent University Salisburylaan 133, B-9820 Merelbeke, Belgium 2. RAVON, Postbus 1413, 6501 BK Nijmegen, The Netherlands 3. Research Institute for Nature and Forest, Kliniekstraat 25, B-1070 Brussels, Belgium 4. Natuurbalans-Limes Divergens, Radboud Universiteit, Mercator III, Postbus 6508, 6503 GA Nijmegen, The Netherlands 5. Department of Obstetrics, Reproduction and Herd Health, Faculty of Veterinary Medicine, Ghent University, Salisburylaan 133, B-9820 Merelbeke, Belgium

EcoHealth, 2013, 10: 344–347

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Abstract Exotic invasive bullfrogs (Lithobates catesbeianus) are considered to exert a considerable negative impact on native amphibian communities. This can be due to competition and predation, but they are also a notorious source of the infectious diseases chytridiomycosis and ranavirosis, affecting amphibian populations globally. Little is known regarding their carriage of other microbial agents that might be transferred to humans or other animals. In this study we determined the occurrence of the amphibian pathogens ranaviruses and Batrachochytrium dendrobatidis and of the zoonotic agents Coxiella burnetii, Neospora caninum, Leptospira sp., Toxoplasma gondii, Mycoplasma sp., Campylobacter sp., Salmonella sp. and broad- spectrum beta-lactamase producing Escherichia coli in 164 bullfrogs from three populations in Belgium and The Netherlands. Although B. dendrobatidis was present at a high prevalence of 63%, mean infection loads were low with an average of 10.9 genomic equivalents (SD 35.5), confirming the role of bullfrogs as B. dendrobatidis carriers, but questioning their role as primary reservoirs for B. dendrobatidis transmission to native amphibian communities. All tested samples were negative for the other infectious agents examined. These results suggest a limited role of bullfrogs as carrier of these pathogens.

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Bullfrogs (Lithobates catesbeianus) have been accidentally introduced in several European countries and have established substantial breeding populations in France, Italy, Germany, Greece, Belgium and The Netherlands (Ficetola et al., 2007; Veenvliet 1996). They are considered to exert a considerable negative impact on native amphibian communities by competition and predation (Johnson et al., 2011; Kiesecker and Blaustein 1998; Kupferberg 1997; Pearl et al., 2004). Additionally, bullfrogs are a notorious source of the notifiable infectious diseases chytridiomycosis and ranavirosis, affecting amphibian populations globally (Garner et al., 2006; Sharifian-Fard et al., 2011). Besides their impact on amphibian populations, bullfrogs could be a source of microbial pathogens for other animals or humans. Bullfrogs in Belgium and The Netherlands are often found in ponds created for recreational fishing and gardens, rendering indirect contact with humans or animals through water likely (Jooris, 2002; Veenvliet, 2009). On several occasions, amphibians have been implicated as a source of Salmonella infections to humans (Chambers and Hulse, 2006; Clarkson et al., 2010; Everard et al., 1979; Sharma et al., 1974; Singh et al., 1979). Leptospira has also been isolated from amphibians, suggesting that amphibians may promote Leptospira infections in aquatic environments (Diesch et al., 1966, 1970; Gravekamp et al., 1991; Everard et al., 1988, 1990). However, little is known regarding amphibians carrying the zoonotic agents Coxiella burnetii, Neospora caninum, Toxoplasma gondii, Mycoplasma sp. or thermotolerant Campylobacter sp. The environment, including wild birds and mammals (Guenther et al., 2011; Garmyn et al., 2011), constitutes an important reservoir of broad-spectrum beta-lactamase producing Escherichia coli isolates. broad-spectrum beta-lactamase are enzymes which are responsible for resistance to Beta-lactam antibiotics like penicillins, cephalosporins and carbapenems. The emergence and wide dissemination of these broad-spectrum beta-lactamases constitute a serious public health concern (Savard and Perl, 2012). Whether amphibians play a role as a reservoir of these strains is not known. We sampled 164 bullfrogs from three populations in Belgium and The Netherlands to evaluate the extent to which these frogs were hosts for the chytrid fungus B. dendrobatidis, ranaviruses and the zoonotic pathogens C. burnettii, N. caninum, Leptospira sp., T. gondii, Mycoplasma sp., Campylobacter sp., Salmonella and of broad-spectrum beta-lactamase producing E. coli. This work will help to determine whether recommendations to prevent the spread of bullfrogs and to eradicate their populations are justified (Doubledee et al., 2003; Ficetola et al., 2007; Govindarajulu et al., 2005; Luja and Rodriguez-Estrella, 2010).

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One hundred and sixty-four clinically healthy L. catesbeianus were collected from 3 populations in Belgium and The Netherlands between June 2010 and October 2011. In Belgium, six animals were collected from a bullfrog population located at a private fishing pond in Hoogstraten (N51_280 E04_450) and 34 animals were collected from a private pond in Arendonk (N51_190 E05_060) (Louette et al., 2012). In The Netherlands, animals were captured from a population in Baarlo (N51_200 E06_050). All animals were euthanized for an invasive species eradication project (INVEXO). This eradication programme for bullfrogs was recently initiated in the framework of the Interreg project ‘Fighting alien invasive species along the Dutch–Belgian border (northwest Europe)’ (INVEXO—Invasieve exoten in Vlaanderen en Zuid-Nederland, Interreg IVa-VLANED- 2.31). Samples of skin, liver, heart, lung, kidney and intestines were taken immediately after euthanasia and stored at -20°C until further use. Intestinal content for bacterial isolation was processed immediately. All faecal samples were analyzed using the ISO 6579:2002 method for the isolation of Salmonella. For the isolation of broad-spectrum beta-lactamase producing E. coli, the intestinal samples were inoculated within 4h onto MacConkey agar plates (Oxoid Ltd., Basingstoke, UK) supplemented with ceftiofur (8 mg/L). After overnight aerobic incubation at 37°C, suspected E. coli colonies were purified on Columbia agar with 5% sheep blood (blood agar, Oxoid) and phenotypically identified. For the isolation of Campylobacter sp., faecal samples were plated on modified charcoal cefoperazone deoxycholate agar (mCCDA, CM0739; Oxoid) supplemented with CCDA selective supplement (SR0155E; Oxoid) and Campylobacter-specific growth supplement (SR0232E; Oxoid), followed by microaerobic incubation at 42°C for 48 h. For the molecular detection of the above-listed pathogens, DNA was extracted from skin, lung, heart, kidney and liver using the DNEasy Tissue Kit (Qiagen, Hilden, Germany). DNA from the intestinal content was prepared using the Stool kit (Qiagen). The DNA from the skin was used in the qPCR to detect B. dendrobatidis (Boyle et al., 2004). All samples were run in duplicate. To control and estimate inhibition, a subset of samples (n = 50) were retested under the same conditions as described above, but with an exogenous internal positive control (VICTM probe, Life technologies, Austin, TX, USA) included as described by Hyatt et al. (2007). We did not find any indications for PCR inhibition. The PCR to detect ranaviruses using the MP4 and 5 primers described by Mao et al., (1997) was performed on the DNA samples obtained from the liver. On the lung samples the PCR described by Van Kuppleveld et al. (1992) was performed to detect Mycoplasma sp. The intestinal derived DNA was used

156 STUDY 6 in PCRs to detect Salmonella sp. (Chiu and Ou, 1996) and Campylobacter sp. (Linton et al., 1996). The detection of T. gondii (used samples: heart and liver), N. caninum (used samples: heart and liver), Leptospira sp. (used samples: liver and kidney) and C. burnetti (used samples: intestines and liver) was done using commercial PCR kits (Adiagene, Paris, France). The absence of a given pathogen was calculated using Win Episcope 2.0. Salmonella, Campylobacter and broad-spectrum beta-lactamase carrying E. coli were not isolated from any of the samples. All samples were negative in the PCRs for the detection of ranaviruses, C. burnetii, N. caninum, Leptospira sp., T. gondii, Mycoplasma sp., Campylobacter sp., Helicobacter sp. and Salmonella. The absence of detection of a given pathogen results in an estimated prevalence of 0% with a maximum prevalence of 39, 8.3 and 2.3% (95% confidence interval) for the three respective populations. One hundred and four out of 164 samples tested positive for B. dendrobatidis. Positive animals were present in the three tested populations. The genomic equivalents (GE) ranged between 1.6 and 368 (mean GE 10.9, SD 35.5) (Table 1).

Table 1. Prevalence of B. dendrobatidis in three populations of invasive bullfrogs in Belgium and The Netherlands.

Population site No. of tested Prevalence of Range of Mean B. dendrobatidis Standard animals B. dendrobatidis (%) GE load (GE) deviation

Hoogstraten (Belgium) 6 67 1.3-4.3 1.6 1.6 Arendonk (Belgium) 34 62 4.4-102 21 30.3 Baarlo (The Netherlands) 124 68 1.6-368 8.6 37.4

The high prevalence (63%) of chytrid infection in adult bullfrogs is comparable with the results found by Garner et al (2006). In their study, the prevalence varied between 0 and 80% in adult bullfrogs in different locations across Europe, Canada and the United States. Since efficient transmission of B. dendrobatidis is promoted both by high infection loads and high prevalence, it is unclear which role bullfrogs play in maintaining B. dendrobatidis in amphibian assemblages. However, high prevalence combined with generally low infection loads, with the exception of erratic Bd-caused mortalities (Pasmans et al., 2010), appear to be the rule rather than the exception in amphibian assemblages under northern European conditions (Garner et al., 2006; Martel et al., 2012), suggesting low infection loads to be sufficient for maintaining B. dendrobatidis. Given the recent discovery of nonamphibian hosts for B. dendrobatidis (McMahon et al., 2013), we hypothesize bullfrogs to be part of a

157 STUDY 6 complex system promoting the persistence of B. dendrobatidis, without adding support to their role as primary contributors to the spread of B. dendrobatidis, as questioned by Liu et al., (2013). Sharifian-Fard et al. (2011) reported a low prevalence (0.75%) of ranaviruses in bullfrog tadpoles. In our study no positive samples for ranaviruses were detected. It is possible that the adults survived a past ranavirus infection and have developed antibodies as was shown for Bufo marinus (Zupanovic et al., 1998). We conclude that invasive bullfrogs in Belgium and The Netherlands do not constitute a significant reservoir for the zoonotic pathogens tested. The relatively low body temperature of the ectothermic anuran host might not provide a suitable environment for these pathogens, mainly occurring in endotherm hosts or, in case of Salmonella, occurring in ectothermic hosts reaching relatively high body temperatures, such as many reptiles.

Acknowledgments This study was partly performed in the framework of EU Interreg IVA project IVA- VLANED-2.31 ‘Invasieve exoten in Vlaanderen en Zuid-Nederland (IN EXO)’.

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In the three following sections, the major findings of this research will be discussed in a broader context, with a brief overview of future research possibilities.

1. Disease situation in Belgian native amphibians 1.1. Chytridiomycosis Study 1 has shown 11 Batrachochytrium dendrobatidis-positive common toads (0.63% prevalence) at nine sites (three in the province of East Flanders, three in Flemish Brabant, and three in Limburg) in Belgium (Sampling localities is shown in Figure 1). The combination of low prevalence, low B. dendrobatidis load in infected animals, absence of obvious negative population trends, and absence of confirmed chytridiomycosis outbreaks, suggests that B. dendrobatidis is endemic at a low intensity in the Flemish region. Spitzen-van der Sluijs et al., (2014) reported that B. dendrobatidis is present in amphibian assemblages in the Netherlands since 1999. Nevertheless, no chytridiomycosis outbreaks are reported except an erratic B. dendrobatidis-caused mortality in Belgium (Pasmans et al., 2010). This endemism of B. dendrobatidis may coincide with environmentally driven B. dendrobatidis prevalence fluctuations that preclude the buildup of B. dendrobatidis infections beyond the critical threshold for large-scale mortality and host population crashes (Spitzen-van der Sluijs et al., 2014). This endemic situation in northern Europe is in contrast with the situation in southern Europe where outbreaks of chytridiomycosis affect midwife toads Alytes obstetricans (Bosch et al., 2001). The differences in outcome of B. dendrobatidis-infections in northern and southern Europe may be associated with a complex interaction between the innate characteristics of individual species, the pathogen and the environment. Depending on those factors, the outcome for a species can range from extinction of the species as the worst-case scenario, to a stable population with sporadic death due to infection (DEH, 2006). Although the sporadic deaths due to B. dendrobatidis infection appear to be a predominant pattern in Belgian native amphibians, the endemism of B. dendrobatidis may result in an on-going pressure especially on small populations of susceptible species facilitating future stochastic events to potentially lead to a local extinction. However, it remains unknown whether the B. dendrobatidis lineage in the Belgian amphibians belongs to the hyper-virulent panzootic line that poses a serious worldwide threat for several amphibian populations. In conclusion, although several knowledge gaps preclude far-reaching conclusions, the impact of chytrid infections caused by B. dendrobatidis in Belgian amphibian assemblages appears negligible given the current conditions. Therefore, we do not recommend the control of B. dendrobatidis to be of primary importance in Belgian amphibian conservation strategies. However, given

165 GENERAL DISCUSSION the importance of environmental determinants in B. dendrobatidis disease epidemiology, a changing world (e.g. climate change) may shift the current state of endemism to a chytridiomycosis epidemic. Putting in place an efficient early warning system and raising public awareness would be an important step to quickly detect changing B. dendrobatidis epidemiology. Limiting the likeliness of introducing new and highly virulent B. dendrobatidis lineages, for example by obligatory absence of B. dendrobatidis in any amphibian traded, would be a second measure that could be implemented relatively easily. While in northern Europe, amphibians appeared to be able to co-exist with B. dendrobatidis, a second chytrid pathogen named Batrachochytrium salamandrivorans has invaded the Netherlands resulting in a disease outbreak and population decline in fire salamanders (Martel et al., 2013). Since this new species was undetectable using previous methods, the challenge is now to define the true risks of B. salamandrivorans either or not in combination with B. dendrobatidis to native amphibians by monitoring of populations and defining the novel fungus host range.

1.2. Ranavirosis Study 1 also revealed that, at present, the prevalence of ranavirus infections in adult common toads (Bufo bufo) is very low (< 0.2% with a confidence interval of 95%) in Flanders (Sampling localities is shown in Figure 1). However, migrating common toads (adults) might not reflect the overall prevalence of ranavirus infections in the toad populations. The reasons for this may be: 1) since only those individuals with a healthy body condition could be expected to participate to spring migration, this may have biased results; 2) since the adults can cope with the infection by developing antibodies (Zupanovic et al., 1998), the absence of an infection in adult populations might not be representative of the absence of the virus within an entire population, including other life stages. Moreover, amphibians can chronically be infected with ranaviruses (Balseiro et al., 2009). If true, ranaviruses may be present in some native amphibian populations below the critical threshold for mortality and therefore, a die-off due to the emergence of ranaviruses can not be excluded in the future. The emergence of ranaviruses in amphibian populations may be promoted by environmental stressors that are natural (e.g., competition, breeding) or human-related (e.g., habitat destruction, poor water quality, climate change). These stressors may directly or indirectly make amphibians more susceptible to infection and lead to outbreaks of viral diseases in amphibian populations. For instance, poor water quality caused by cattle activity around wetlands was linked to increased ranaviruses prevalence in pond-breeding amphibians (Gray et al., 2007; Hoverman et al.,

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2012). Thus, cattle ponds and run-off retention ponds can be hotspot of ranaviruses disease due to enhanced environmental stress. Therefore, expanding this study to include additional species and monitoring several life stages will be helpful in further characterizing the epidemiology of ranaviruses in native amphibians. In conclusion, ranaviruses are present in Belgium and more or less continuous introduction of novel viral strains from neighbouring countries, amphibian trade and translocations is highly likely (please see further). Since ranavirosis in wild amphibians has hitherto not been reported from Belgium, we would propose the same actions to be taken as proposed for B. dendrobatidis, i.e. raising public awareness, an early warning system and amphibian trade restrictions. These measures could thus be implemented for both notifiable diseases and require a minimal investment of the regional governments. This would be a unique opportunity and a first step to expand “wildlife population health” beyond the current, highly anthropocentric approach (where wildlife disease is currently only deemed of interest if directly relevant to human and/or livestock health), and truly invest in controlling wildlife diseases to limit their impact on biodiversity.

2. Disease threats to Belgian native amphibians The following paragraphs explain pathogen introduction pathways that can pose a threat to our endangered amphibian species.

2.1. Introduction of amphibian pathogens by invasive species The American bullfrog (Lithobates catesbeianus) is listed as one of the “100 of the world’s worst invasive alien species” (Lowe et al., 2000). They can pose serious threats to native species and play an important role in the deterioration of amphibian populations by transmitting infectious diseases agents such as B. dendrobatidis, ranaviruses and Chlamydiales (Daszak et al., 2004; Hanselmann et al., 2004; Garner et al., 2006; Martel et al., 2012). Although outbreaks of ranaviruses with high mortality rates have been described in ranaculture facilities, bullfrogs are considered to be ranavirus reservoirs based on prevalence and viral load (Miller et al., 2007; Mazzoni et al., 2009; Une et al., 2009). This difference in ranaviral-infection outcome in American bullfrog populations may be associated with environmental factors, genetic diversity and stress (Gahl and Calhoun, 2008; Ridenhour and Storfer, 2008). FV3 and FV3-like ranaviruses have been detected in American bullfrogs following an outbreak at an amphibian farm in Brazil (Mazzoni et al., 2009).

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Figure 1. Sampling localities of common toads (Bufo bufo) and American bullfrogs (Lithobates catesbeianus) in Flanders (Belgium) and The Netherlands.

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We detected the common midwife toad virus (CMTV) in clinically healthy bullfrog tadpoles in Belgium (Study 2) showing that the bullfrog in Flanders, as is the case in the United States of America (Adams & Pearl, 2007), can act as a reservoir for ranaviruses. Although ranavirus infection in this species was detected at a low prevalence (0.75%) and is probably not a major cause of mortality in wild populations, there is a potential threat of spill over to naïve indigenous amphibians. American bullfrogs are also known carriers of B. dendrobatidis (Garner et al., 2006; Gray et al., 2007). In 2008 and in 2009, 88 samples of American bullfrogs were collected in Flanders. The result showed that 18 samples were B. dendrobatidis-positive (20.5% prevalence) (Spitzen-van der Sluijs et al., 2010). We reported that B. dendrobatidis is highly prevalent in two populations of invasive bullfrogs in Belgium (67% and 62% prevalence in Hoogstraten and Arendonk, respectively, study 6, Figure 1). Since they are thought to be tolerant to B. dendrobatidis, bullfrogs can move the pathogen to new locations and expose susceptible amphibians. The actual contribution of bullfrogs to dissemination of B. dendrobatidis remains unclear, since the fungus can also be transmitted by waterfowl (Garmyn et al., 2012) or crayfish (Procambarus spp.) (McMahon et al., 2013). Moreover, we identified a novel species of Chlamydiales (Candidatus Amphibiichlamydia ranarum), with a prevalence of 71% (N = 200), in bullfrog tadpoles from the Baarlo population in the Netherlands (study 3, Figure 1). Although none of the examined tadpoles showed signs of clinical disease, the pathogenic potential for native amphibian species should be investigated since the other novel Amphibiichlamydia named Candidatus Amphibiichlamydia salamandrae has been associated with mortality in captive salamanders (Martel et al., 2013). While we know little regarding drivers of emergence of Candidatus Amphibiichlamydia in Belgium, changes in the host–environment and the disease ecology can be key to creating novel transmission patterns and selection of novel pathogens with fitter genetic traits (Engering et al., 2013). Thus, understanding the dynamics of these novel pathogens in amphibian populations is crucial to prevent amphibian declines.

2.2. Translocation of pathogens through commercial trade of amphibians Commercial trade in amphibians such as the commercial exploitation of larval tiger salamanders (Ambystoma tigrinum) as fishing bait in the United States of America or African clawed frog (Xenopus laevis) for research and as pets provides a vehicle for the global movement of amphibian hosts and their associated pathogens (Picco and Collins, 2008). B. dendrobatidis has been detected in amphibians traded as pets, food and biomedical research

169 GENERAL DISCUSSION organisms from different countries (Cunningham et al., 2003). Similarly, ranaviruses can be translocated over large distances as a result of international commerce of their vertebrate hosts i.e. amphibians, reptiles and bony fish (Lesbarrères et al., 2011). For example, ranaviruses associated with an amphibian die-off in the UK were more similar to North American ranaviruses than they were to ranaviruses from other localities, including continental Europe, which suggests anthropogenic introduction (Hyatt et al., 2000). Such introductions may be also of concern in Belgium, as a study reported ranavirus-associated mass mortality in imported red-tailed knobby newts (Tylototriton kweichowensis) in Belgium (Pasmans et al., 2008). It is clear that without effective implementation of national and international bio- security measures, the occurrence, trans-boundary spread and impact of emerging infectious diseases will continue putting pressure on native amphibian populations.

2.3. Introduction of pathogens from outbreaks in neighboring countries Although the primary pathogen introduction pathway is the amphibian trade, a secondary pathway is considered to be as a consequence of natural dispersal of the hosts of pathogens from neighbouring countries (Gozlan et al., 2010). Therefore, further expansion of pathogens towards Belgian amphibians can be expected from neighbouring countries where disease outbreaks have been recorded. In the past four years, ranaviruses have resulted in mass mortalities in green frog populations in the Netherlands and Germany. In 2010, an outbreak of common midwife toad virus (CMTV) resulted in a mass die-off of over 1,000 wild water frogs (Pelophylax spp.) and 10 smooth newts (Lissotriton vulgaris) in the Netherlands (Kik et al., 2010). Consequently, transmission of infection from this neighbouring country is considered as a potential threat to Belgian amphibians. Moreover, Stöhr et al. (2013) reported that ranaviruses are responsible for a die-off of several edible frogs (Pelophylax kl. esculentus) in Germany. Thus, broad-scale monitoring for ranavirus infections, implementation of control measures, and education of the public to prevent the disease from spreading of the pathogen from neighbouring countries is warranted.

2.4. Introduction of pathogens by live feed items Feed items used as fish bait can also be infected with pathogens and posed a potential threat to native amphibian communities. Study 4 revealed that the role of chironomid larvae (bloodworms) that are widely disseminated in the environment as fish bait pose a minor risk of spreading pathogens i.e. ranaviruses and B. dendrobatidis to native amphibians. Although

170 GENERAL DISCUSSION there may be a minor risk for transmission of infectious agents, the excessive levels of Pb in the bloodworms revealed the potential threat of Pb accumulation in native amphibians.

3. The zoonotic risk of handling amphibians Different serotypes of Salmonella have been isolated from amphibians. While these isolates have frequently been reported from (sub) tropical regions (O’Shea et al., 1990; Drake et al., 2012), little is known regarding the role of wild amphibians to maintain Salmonella infection cycles in temperate regions. Study 5 showed that migrating common toads living in Belgium probably play a negligible role in maintaining Salmonella infection cycles. Moreover, we reported (study 6, Figure 1) that all 164 bullfrog samples from Belgium and the Netherlands tested negative for the following zoonotic agents: Coxiella burnetii, Neospora caninum, Leptospira sp., Toxoplasma gondii, Mycoplasma sp., Campylobacter sp., Salmonella sp., and extended-spectrum, beta-lactamase-producing Escherichia coli. The reason for the absence of these zoonotic agents can be associated with the thermal niche of amphibians in temperate regions. In particular, Salmonella growth is substantially reduced at temperatures below 10°C. The relatively low body temperature of the ectothermic anuran host might not provide a suitable environment for these pathogens. The results of our studies (5&6) support the hypothesis that the significance of ectothermic vertebrates as Salmonella reservoirs is limited to those species that provide sufficiently high body temperatures for successful, persistent host colonisation (Pasmans et al., 2013). However, seasonal variations in Salmonella infection have been reported, with both their prevalence and severity being greater during the warmer months (Haley et al., 2009). Therefore temperature is considered a likely factor influencing Salmonella occurrence in animal hosts. The occurrence of Salmonella infections in amphibians may thus be higher during summer months. The absence of zoonotic agents in common toads and bullfrogs indicated that the prevalence of those agents in amphibian populations is currently low in Flanders and of low public health concern. In conclusion, our studies (5&6) provided evidence that handling amphibians in northern European countries with temperate climates poses no or a minor zoonotic risk. However, a potential seasonal fluctuation in pathogen prevalence requires more research to be conducted in a broader variety of wild amphibians during the warmer months.

171 GENERAL DISCUSSION

References Adams MJ, Pearl CA. 2007. Problems and opportunities managing invasive Bullfrogs: is there any hope? Biological Invaders in Inland Waters: Profiles, Distribution and Threats: 679-693. Baslserio A, Dalton KP, delCerro A, Márquez I, Parra F, Prieto JM, Casais R. 2009. Outbreak of common midwife toad virus in alpine newts (Mesotriton alpestris cyreni) and common midwife toads (Alytes obstetricans) in Northern Spain: A comparative pathological study of an emerging ranavirus The Veterinary Journal 186(2):256-8. Bertsch P.M, Janecek L, Rosier B. 2001. Technical Report: Savannah River Ecology Laboratory Annual Technical Progress Report of Ecological Research. Pp 83. Blaustein AR, Wake DB. 1990. Declining amphibian populations: a global phenomenon? Trends in Ecology and Evolution 5:203-4. Bosch J, Martínez-Solano I, García-Paris M. 2001. Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricans) in protected areas of central Spain. Biological Conservation 97(3):331-7. Cunningham AA, Daszak P, Rodriguez JP. 2003. Pathogen pollution: defining a parasitological threat to biodiversity conservation. Journal of Parasitology 89:78-83. DEH. 2006. Infection of amphibians with chytrid fungus resulting in chytridiomycosis. Department of the Environment and Heritage. 29 pp. http://www.environment.gov.au/system/files/resources/8d01e983-3619-4d83-9b5a- 6f9fb4d34e3b/files/chytrid-report.pdf. Daszak P, Strieby A, Cunningham AA, Longcore JE, Brown CC, Porter D. 2004. Experimental evidence that the bullfrog (Lithobates catesbeianus) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians. Herpetological Journal 14(4):201-7. Drake M, Amadi V, Zieger U, Johnson R, Hariharan H. 2012. Prevalence of Salmonella spp. in cane toads (Bufo marinus) from Grenada, West Indies, and their antimicrobial susceptibility. Zoonoses and Public Health 60(6):437-41. Engering A , Hogerwerf L, Slingenbergh J. 2013. Pathogen–host–environment interplay and disease emergence. Emerging Microbes and Infections 2:2, e5. Gahl MJ, Calhoun AJK. 2008. Landscape setting and risk of Rnavirus mortality events. Biology Conservation141:2679–89.

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Garmyn A, Van Rooij P, Pasmans F, Hellebuyck T, Van Den Broeck W, Haesebrouck F, Martel A. 2012. Waterfowl: potential environmental reservoirs of the chytrid fungus Batrachochytrium dendrobatidis. PloS ONE 7(4): e35038. Garner TWJ, Perkins MW, Govindarajulu P, Seglie D, Walker S, Cunningham AA, Fisher MC. 2006. The emerging amphibian pathogen Batrachochytrium dendrobatidis globally infects introduced populations of the North American bullfrog, Lithobates catesbeianus. Biology Letters 2(3):455–9. Gozlan RE, Andreou D, Asaeda T, Beyer K, Bouhadad R, Burnard D, Caiola N, Cakic P, Djikanovic V, Esmaeili H.R, Falka I, Golicher D, Harka Á, Jeney G, Kovac V, Musil J, Nocita A, Porz M, Poulet N, Virbickas T, Wolter C, Tarkan SA, Tricarico E, Trichkova T, Verreycken H, Witkowski A, Zhang CG, Zweismueller I, Britton RJ. 2010. Pancontinental invasion of Pseudorasbora parva: towards a better understanding of freshwater fish invasions. Fish and Fisheries 11: 315–340. Gray MJ, Miller DL, Schmutzer AC, Baldwin CA. 2007. Frog Virus 3 prevalence in tadpole populations at cattle-access and non-access wetlands in Tennessee, USA. Diseases of Aquatic Organisms 77:97–103. Haley BJ, Cole DJ, Lipp EK. 2009. Distribution, diversity, and seasonality of waterborne salmonellae in a rural watershed. Applied and Environmental Microbiology 75(5): 1248-55. Hanselmann R, Rodríguez A, Lampo M, Fajardo-Ramos L, Aguirre AA, Kilpatrick AM, Rodríguez J, Daszak P. 2004. Presence of an emerging pathogen of amphibians in introduced bullfrogs Lithobates catesbeianus in Venezuela. Biological Conservation 120(1):115-9. Hoverman JT, Gray MJ, Miller DL, Haislip NA. 2012.Widespread Occurrence of Ranavirus in Pond-Breeding Amphibian Populations. EcoHealth 9(2):36–48. Hyatt AD, Gould AR, Zupanovic Z, Cunningham AA, Hengstberger S, Whittington RJ, Kattenbelt J, Coupar BEH. 2000. Comparative studies of piscine and amphibian iridoviruses. Archives of Virology 145(2):301-331. Kik MMA, Spitzen-van der Sluijs A, Pasmans F, Wohlsein P, Gröne A, Rijks JM. 2011. Ranavirus-associated mass mortality in wild amphibians, the Netherlands, 2010: A First Report. The Veterinary Journal 190(2):284-6. Lesbarrères D, Balseiro A, Brunner J, Chinchar VG,Duffus A, Kerby J, Miller DL, Robert J, Schock DM, Waltzek T, Gray MJ. 2011. Ranavirus: past, present and future. Biology Letters 8:481−483.

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Lowe S, Browne M, Boudjelas S. 2000. 100 of the World's Worst Invasive Alien Species. Auckland, New Zealand: IUCN/SSC Invasive Species Specialist Group. 11 pp. http://www.issg.org/database/species/reference_files/100English.pdf. Martel A , Adriaensen C, Bogaerts S, Ducatelle R, Favoreel H, Crameri S, Hyatt AD, Haesebrouck F, Pasmans F. 2012. Novel Chlamydiaceae disease in captive salamanders. Emerging Infectious Diseases 18(6). Martel A, Spitzen-van der Sluijs A, Blooi M, Bert W, Ducatelle R, Fisher MC, Woeltjes A, Bosman W, Chiers K, Bossuyt F, Pasmans F. 2013. Batrachochytrium salamandrivorans sp. nov. causes lethal chytridiomycosis in amphibians. Proceedings of the National Academy of Sciences USA 110(38):15325-9. Mazzoni R, José de Mesquita A, Fleury LFF, Diederichsen de Brito WME, Nunes IA, Robert J, Morales H, Coelho ASG, Barthasson DL, Galli L, et al. 2009. Mass mortality associated with a frog virus 3-like Ranavirus infection in farmed tadpoles, Lithobates catesbeianus, from Brazil. Diseases of Aquatic Organisms 86:181–91. McMahon TA, Brannelly LA, Chatfield MWH, Johnson PTJ, Joseph MB, McKenzie VJ, Richards-Zawacki CL, Venesky MD, Rohr JR. 2013. Chytrid fungus Batrachochytrium dendrobatidis has nonamphibian hosts and releases chemicals that cause pathology in the absence of infection. Proceedings of the National Academy of Sciences USA 110(1): 210-215. Meyer AH, Schimidt BR, Grossenbacher K.. 1998. Analysis of three amphibian populations with quarter-century long time-series. Proceedings of the Royal Society B: Biological Sciences 265(1395):523-8. Miller DL, Rajeev S, Gray MJ, Baldwin C. 2007. Frog Virus 3 infection, cultured American bullfrogs. Emerging Infectious Diseases 13:342–3. O’shea P, Speare R, Thomas AD. 1990. Salmonella serotypes from the cane toad, Bufo marinus. Australian Veterinary Journal 67:310. Pasmans F, Blahak S, Martel A, Pantchev N, Zwart P. 2008. Ranavirus associated mass mortality in imported red tailed knobby newts (Tylototriton kweichowensis): A case report. The Veterinary Journal 176:257–259. Pasmans F, Muijsers M, Maes S, Van Rooij P, Brutyn M, Ducatelle R, Haesebrouck F, Martel A. 2010. Chytridiomycosis related mortality in a midwife toad (Alytes obstetricans) in Belgium. Vlaams Diergeneeskundig Tijdschrift 79:461-3.

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Pasmans F, Boyen F, Haesebrouck F. 2013. Salmonella in exotic pet animals. In: Barrow, P.A. (ed.) Salmonella in domestic animals. CABI publishing, Oxfordshire, UK. Pp:337-50. Picco AM, Collins JP. 2008. Amphibian commerce as a likely source of pathogen pollution. Conservation Biology 22(6):1582–1589. Ridenhour BJ, Storfer AT. 2008. Geographically variable selection in Ambystoma tigrinum virus (Iridoviridae) throughout the western USA. Journal of Evolutionary Biology 21(4):1151-9. Spitzen-Van der Sluijs AM, Zollinger R, Bosman W, Van Rooij P, Clare F, Martel A, Pasmans F. 2010. Short Report Batrachochytrium dendrobatidis in amphibians in the Netherlands and Flanders (Belgium). Stichting RAVON Report 2009-29, Nijmegen. 32 pp. Spitzen – van der Sluijs A, Martel A, Hallmann CA, Bosman W, Garner TWJ5, Van Rooij P, Jooris R, Haesebrouck F, Pasmans F. 2014. Environmental determinants of recent endemism of Batrachochytrium dendrobatidis infections in amphibian assemblages in the absence of disease outbreaks. Conservation Biology DOI: 10.1111/cobi.12281.(early online edition). Stöhr AC, Hoffman A, Papp T, Robert N, Pruvost NBM, Reyer HU, Marschang RE. 2013. Long-term study of an infection with Ranavirus in a group of edible frogs and partial characterization of two viruses based on four genomic regions. The Veterinary Journal 197:238-244. Stuart SN. 2012. Responding to the amphibian crisis: too little, too late? Alytes 29(1-4): 9-12. Une Y, Sakuma A, Matsueda H, Nakai K, Murakami M. 2009. Ranavirus outbreak in North American bullfrogs (Lithobates catesbeianus), Japan. Emerging Infectious Diseases 15:1146–7. Zupanovic Z, Lopez G, Hyatt AE, Green B, Bartran G, Parkes H, Whittington RJ, Speare R. 1998. Giant toads Bufo marinus in Australia and Venezuela have antibodies against ‘ranaviruses’. Diseases of A uatic Organisms 32: 1–8.

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176

SUMMARY

176 173 SUMMARY

178 SUMMARY

Amphibian populations are at the forefront of the global biodiversity crisis, with more extinct or declining species than any other class of vertebrates. Of the fifteen Belgian amphibian species, 10 are decreasing in population and currently threatened. Although putative causes of observed declines are numerous and frequently controversial, the emergence of infectious diseases such as chytridiomycosis and ranavirosis, and invasive species such as American bullfrogs (Lithobates catesbeianus), represent two of the greatest threats confronting native amphibians. One of the main issues hampering amphibian conservation is the limited knowledge with regard to amphibian infections, which is currently heavily focused on chytridiomycosis and ranavirosis. The aim of this PhD study was to gain insight in microbiological infections in native amphibians, which are of importance in amphibian conservation related issues. In the first part of this work, we determined the presence of bacteria, viruses and fungi in native and invasive amphibians with the potential to affect amphibian health. In Study 1, road-killed common toads (Bufo bufo; n=1740) from Flanders, Belgium, were tested for Ranavirus and Batrachochytrium dendrobatidis using polymerase chain reaction. Both infections were present at a very low prevalence (<0.2% with a confidence interval of 95% for Ranavirus and 0.63% for B. dendrobatidis). The combination of low prevalence, low B. dendrobatidis load in infected animals, absence of obvious negative population trends, and absence of confirmed chytridiomycosis outbreaks, suggests that B. dendrobatidis is endemic at a low intensity in the Flemish region. In Study 2, the role of bullfrogs as carriers of ranaviruses in Europe was assessed. For this purpose, we collected 400 clinically healthy tadpoles of L. catesbeianus from 3 invasive bullfrog populations at Hoogstraten, Belgium (51°47′Ν, 4°75′Ε) during May–June 2010. A low prevalence of Ranavirus infection (0.75%) in the bullfrog tadpoles was examined. This study shows that invasive bullfrogs, a known reservoir of chytridiomycosis, are also a likely carrier of ranaviral disease in Europe. A novel species of Chlamydiales (Candidatus Amphibiichlamydia ranarum) was identified with a prevalence of 71% in bullfrog tadpoles from an introduced population in the Netherlands in Study 3. Although none of the tadpoles examined showed signs of clinical disease, urgent evaluation of its pathogenic potential for native amphibian species is required since its closest relative ‘Candidatus Amphibiichlamydia salamandrae’ has been associated with severe disease and high mortality in captive salamanders. In the second part of this work (study 4) we tried to estimate the risk of providing live feed items to amphibians in ex situ programmes. The global amphibian decline phenomenon has rendered ex situ conservation a currently widely accepted measure to preserve amphibian

179 SUMMARY biodiversity. However, the predatory nature of most amphibians requires the feeding of live and often wild caught feed items that may pose a risk to amphibian health and thus ex situ conservation. In this study, we defined the risk of a commonly used food item (Chironomidae or so-called bloodworms) for amphibian health. We found neither zoonotic bacteria nor the amphibian pathogens Ranavirus and B. dendrobatidis in these samples, but excess levels of lead in 2 samples and low content of zinc in 4 samples implicated potential risk of lead accumulation and zinc inadequacy. Proximate nutritional analysis revealed that, although the high protein content in these samples could fulfill the protein needs, the wide range in lipid: protein ratio warrants further study, since this ratio will affect the amount and pathway of energy supply to the amphibians. Our study indicated that, although environmentally- collected chironomids larvae may not be vectors of specific pathogens, they can be associated with nutritional imbalances and may also result in lead bioaccumulation and zinc inadequacy in amphibians. Chironomidae larvae may thus not be recommended as single diet item for amphibians. In the third part, the zoonotic risk of human contact with amphibians during conservation activities such as safeguarding activities for native amphibians and controlling invasive amphibian species was examined. For this purpose, we examined the presence of Salmonella sp. in native common toads (Bufo bufo) (Study 5). Between February and April 2011, the intestinal content of 1740 samples of road killed migrating common toads in five Flemish provinces of Belgium was examined for the presence of Salmonella using bacterial culture and PCR. All the samples were negative. These results suggest that the role of migrating common toads in maintaining the infection cycle of Salmonella in northern European temperate regions is negligible. In study 6 we determined the occurrence of zoonotic agents, i.e. Coxiella burnetii, Neospora caninum, Leptospira sp., Toxoplasma gondii, Mycoplasma sp., Campylobacter sp., Salmonella sp. and broad-spectrum beta-lactamase producing Escherichia coli in invasive American bullfrogs in Belgium. All tested samples were negative for the infectious agents examined. These results suggest a limited role of bullfrogs as carrier of zoonotic pathogens.

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179 SAMENVATTING

182 SAMENVATTING

Amfibieënpopulaties staan in de frontlinie van een algemene biodiversiteitscrisis, met meer uitgestorven soorten en afname in soortenaantallen dan enig andere groep binnen de gewervelden. Van de 15 Belgische amfibieënsoorten, kennen 10 soorten een sterke afname in aantallen en zijn heden in bepaalde mate in hun voortbestaan bedreigd. Hoewel de vermeende oorzaken van deze waargenomen achteruitgang talrijk zijn en vaak controversieel, vormen nieuwe infectieziekten zoals chytridiomycose en ranavirose, én invasieve soorten zoals de stierkikker (Lithobates catesbeianus), momenteel de twee grootste bedreigingen voor inheemse amfibieën. De kennis van infectieziekten bij inheemse amfibieën is zeer beperkt en in hoofdzaak gelimiteerd tot chytridiomycose en ranavirose, wat het nemen van aangepaste behouds-en beheermaatregelen voor amfibieën aanzienlijk bemoeilijkt. Het doel van dit doctoraatsonderzoek is dan ook inzicht te verwerven in microbiële infecties die voorkomen bij inheemse amfibieën en potentieel een bedreiging vormen. In een eerste deel van het doctoraatsonderzoek werden zowel inheemse als invasieve soorten onderzocht op aanwezigheid van bacteriën, virussen en schimmels die mogelijk een negatieve impact hebben op de gezondheid van amfibieën. In een eerste studie werden gewone padden (Bufo bufo; n=1740) die tijdens de voorjaarstrek doodgereden werden in Vlaanderen ingezameld en getest op aanwezigheid van Ranavirus en Batrachochytrium dendrobatidis met behulp van polymerase chain reaction (PCR). Beide infecties kennen een zeer lage prevalentie (<0.2% met een 95% betrouwbaarheidsinterval voor Ranavirus en 0.63% voor B. dendrobatidis). Deze lage prevalentie, in combinatie met een lage infectiegraad van B. dendrobatidis bij geïnfecteerde dieren, afwezigheid van een duidelijke negatieve populatietrend en afwezigheid van bevestigde uitbraken van chytridiomycose, doen vermoeden dat B. dendrobatidis endemisch is in Vlaanderen met een lage infectieintensiteit. In een tweede studie, werd de rol van stierkikkers als drager van Ranavirussen binnen Europa nagegaan. Gedurende de periode mei-juni 2010 werden vierhonderd klinisch gezonde L. catesbeianus larven van 3 stierkikkerpopulaties uit Hoogstraten (België; 51°47′Ν, 4°75′Ε) ingezameld. Bij de onderzochte stierkikkerlarven werd Ranavirus aangetroffen aan een lage prevalentie (0.75%). Dit onderzoek toont aan dat de invasieve stierkikker niet alleen een gekend reservoir is voor chytridiomycose, maar waarschijnlijk ook voor Ranavirus en dus de verspreiding van deze virale ziekte binnen Europa kan bevorderen. In een derde studie werd een nieuwe Chlamydiales soort (Candidatus Amphibiichlamydia ranarum) ontdekt bij stierkikkerlarven in een geïntroduceerde Nederlandse populatie, met een prevalentie van 71%. Het meest aanverwante taxon ‘Candidatus Amphibiichlamydia salamandrae’ werd reeds in

183 SAMENVATTING verband gebracht met ernstige ziekte en hoge sterfte bij salamanders in gevangenschap. Hoewel geen enkele van deze onderzochte larven een klinisch ziektebeeld vertoonden, dient het pathogeen potentieel van deze nieuwe kiem dringend te worden onderzocht. In een tweede deel van het doctoraatsonderzoek (studie 4) werd een risicoanalyse uitgevoerd op het voederen van amfibieën in ex situ behoud- en kweekprogramma’s met levende rode muggenlarven, een standaard voeder. Door de algemene afname in soortaantallen is het behoud van amfibieënpopulaties ex situ een algemeen aanvaarde maatregel om de instandhouding van de biologische diversiteit van amfibieën te vrijwaren. Omdat alle gemetamorfoseerde amfibieën predatoren zijn, moeten deze dieren in gevangenschap gevoederd worden met, meestal levende prooidieren. De kwaliteit van dit voeder kan echter nefast kan zijn voor de gezondheid van amfibieën en dus het ex situ behoud van deze dieren. In deze studie evalueerden we het gezondheidsrisico voor amfibieën van vaak gebruikt levend voeder (Chironomidae of rode muggenlarven). In de voederstalen werden geen zoönotische bacteriën noch de amfibieënpathogenen Ranavirus en B. dendrobatidis teruggevonden. In twee stalen werd echter een loodconcentratie boven de norm waargenomen en in 4 stalen een lage zinkconcentratie, wat aanleiding kan geven tot respectievelijk loodaccumulatie en zinktekort. Voedingsanalyse van de voederstalen toont aan dat het hoge proteïnegehalte van dit voeder kan voldoen aan de proteïnebehoefte van amfibieën. De grote verschillen in verhouding vetten/proteïnen vereisen verder onderzoek, daar deze verhouding de hoeveelheid en hoedanigheid van energievoorziening voor amfibieën beïnvloedt. Dit onderzoek toont aan dat rode muggenlarven geen vectoren zijn van specifieke pathogenen. Het voederen van amfibieën met deze larven kan echter wel aanleiding geven tot een nutritioneel onevenwicht, bioaccumulatie van lood en zinktekort. Rode muggenlarven als enige voedselbron zijn dus niet aangewezen als dieet voor amfibieën. In een derde deel van het doctoraatsonderzoek werd het zoönotisch risico nagegaan voor mensen die in nauw contact komen met amfibieën tijdens natuurbehoud gerelateerde activiteiten, zoals bij monitoring van inheemse soorten of controle op invasieve soorten (Studie 5). Hiervoor werd eerst de aanwezigheid van Salmonella sp. nagegaan in de inheemse gewone pad (Bufo bufo). In 2011 werden tijdens de paddentrek (februari tot april) 1740 doodgereden gewone padden ingezameld in 5 Vlaamse provincies. Vervolgens werd de darminhoud onderzocht op aanwezigheid van Salmonella aan de hand van bacteriecultuur en PCR. Alle stalen testten negatief voor Salmonella. Deze resultaten suggereren dat in de Noord-Europese gematigde gebieden migrerende gewone padden een verwaarloosbare rol spelen in de instandhouding van de infectiecyclus van Salmonella. In een zesde studie werd

184 SAMENVATTING het voorkomen van de zoönotische agentia Coxiella burnetii, Neospora caninum, Leptospira sp., Toxoplasma gondii, Mycoplasma sp., Campylobacter sp., Salmonella sp. en breed- spectrum beta-lactamase producerende Escherichia coli nagegaan bij invasieve stierkikkers in België. Alle stalen testten negatief voor de onderzochte infectieuze agentia. Deze resultaten duiden op een beperkte rol van de stierkikker als drager van zoönotische kiemen.

185 SAMENVATTING

186

CURRICULUM VITAE

181 CURRICULUME VITAE

188 BIBLIOGRAPHY

Mojdeh Sharifian Fard was born on 2 April 1981 in Iran. After graduating from Professor Hesabi School in Tehran in 1999, she started veterinary medicine studies at the University of Tehran. In 2006, she obtained her general veterinarian doctorate. Her thesis was entitled “Prevention of shedding and re-shedding of Toxoplasma gondii oocysts in experimentally infected cats treated with oral clindamycin”. After graduation, she opened a small animal clinic in Tehran and spent three years there. In October 2010, she left Iran to work in the Department of Pathology, Bacteriology and Poultry Diseases, Faculty of Veterinary Medicine, Ghent University. Her doctoral research study was performed under the supervision of Prof. dr. A. Martel and Prof. dr. F. Pasmans and she spent four years researching disease threats to Belgian amphibians. Her research led to several scientific publications in international journals.

189 BIBLIOGRAPHY

190

BIBLIOGRAPHY

190

BIBLIOGRAPHY

192 BIBLIOGRAPHY

1. Publication Sharifian Fard M, Pasmans F, Martel A. 2014. Migrating common toads (Bufo bufo) in rural temperate regions: reservoirs of Salmonella? Journal of Wildlife Diseases 50(2): 326-9. Sharifian Fard M., Pasmans F, Adriaensen C, Du Laing G, Janssens G, Martel A. 2014. Chironomidae bloodworms larvae as aquatic amphibian food. Zoo Biology 33(3): 221– 227. Maazi N, Malmasi A, Shayan P, Nassiri SM, Zahraei Salehi T, Sharifian Fard M. 2014. Molecular and serological detection of Ehrlichia canis in naturally exposed dogs in Iran: An analysis of associated risk factors. Brazilian Journal of Veterinary Parasitology 23(1):16-22. Martel A, Sharifian Fard M, Pasmans F, Adriaensen C. 2013. Absence of zoonotic agents in invasive bullfrogs (Lithobates catesbeianus) in Belgium and The Netherlands. Ecohealth 10(4):344-7. Martel A, Adriaensen C, Sharifian Fard M, Vandewoestyne M, Deforce D, Favoreel H, Bergen K, Spitzen-van der Sluijs A, Devisscher S, Adriaens T, Louette G, Baert K, Hyatt A, Crameri S, Haesebrouck F, Pasmans F. 2013. The novel ‘Candidatus Amphibiichlamydia ranarum’ is highly prevalent in invasive exotic bullfrogs (Lithobates catesbeianus). Environmental Microbiology Reports 5(1):105–8. Malmasi A, Mosallanejad B, Mohebali M, Taheri, Sharifian Fard M. 2013. Effects of topical chloramphenicol and ciprofloxacin on conjunctival bacterial flora of healthy dogs. Clinical Comparative Pathology: Submitted. Martel A, Sharifian Fard M, Van Rooij P, Jooris R, Boone F, Haesebrouck F, Rooij D, Pasmans F . 2012. Road-killed common toads (Bufo bufo) in Flanders (Belgium) reveal low prevalence of ranaviruses and Batrachochytrium dendrobatidis. Journal of Wildlife Diseases 48(3):835–9. Sharifian Fard M, Pasmans F, Adriaensen C, Devisscher S, Adriaens T, Louette G, Martel A. 2011. Ranavirosis in invasive bullfrogs, Belgium. Emerging Infectious Diseases 17(12): 2371-2. Shadkhast M, Shabazkia HR, Bigham-Sadegh A, Ebrahim Shariati SE, Mahmoudi T, Shariffian Fard M. 2010. The morphological characterization of the blood cells in the central Asian tortoise (Testudo horsfieldii). Veterinary Research Forum 1(3):134-41. Malmasi A, Mosallanejad B, Mohebali M, Sharifian Fard M, Taheri M. 2009. Prevention of shedding and re-shedding of Toxoplasma gondii oocytes in experimentally infected cats treated with oral Clindamycin: a preliminary study. Zoonoses and Public Health

193 BIBLIOGRAPHY

56(2):102-4.

2. Conference papers Ghiasi F, Mirzargar S, Sharifian Fard M. 2007. Cadmium exposure of common carp: effect on phagocytic index. 2th European congress of Ichthyology. Shadkhast M, Chavoshi K, Sabouri MH, Sharifian Fard M. 2003. Myocardial bridge in animals. First joint meeting of EACA and AACA, GRAZ.

194

ACKNOWLEDGEMENTS

189 ACKNOWLEDGEMENTS

196 ACKNOWLEDGEMENTS

Firstly, I wish to thank my parents for being so understanding of my time and dedication. I am extremely fortunate to have such a wonderful family. Undoubtedly, I would not be where I am without the lifelong support and encouragement from my father and I cannot thank him enough. I also would not be in this position without the support and guidance of my promoters, Prof. dr. An Martel and Prof. dr. Frank Pasmans. I would like to express my sincere gratitude to them for the continuous support of my Ph.D study and research, for their patience, motivation, enthusiasm, and immense knowledge. Their guidance helped me in all the time of research and writing of this thesis. I thank them for helping to develop a research problem, and then for providing their advice, support and guidance throughout my doctoral programme. The ability to pursue my Ph.D would not have been possible without my department head and dissertation committee chairman, Prof. dr. Freddy Haesebrouck. He has been supportive of my research and has taken every opportunity to allow me to succeed. I would also like to acknowledge my committee members, Prof. dr. Lieven De Zutter, Prof. dr. Hans Nauwynck, Prof. dr. Jeroen Dewulf, Dr. Gerald Louette and Drs. Pascale Van Rooij who graciously agreed to serve on my committee, when they were probably already swamped with work. I would like to thank my friends and colleagues Pascale Van Rooij and Marc Verlinden, who created such a nice atmosphere in the office and shared in unforgettable memories. Thank you and the best of luck in your future endeavours. I would also like to thank Connie Adriaensen, Mark Blooi and Bart Dewulf for generously sharing their expertise and for their helpful suggestions whenever I had problems during the study. I would also like to thank the secretarial staff, Gunter Massaer and Jo Dewaele, for answering all my questions. I extend my sincerest thanks to all

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ACKNOWLEDGEMENTS members of the Department of Bacteriology and all those who contributed directly or indirectly to the dissertation. In particular, I wish to thank Serge Verbanck and Arlette Van de Kerckhove, who taught me the necessary techniques and for their assistance in the implementation of experimental protocols. The work presented in this dissertation would not have been possible without the help of numerous people who collected samples.

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