Universidade de Aveiro Departamento de Biologia

2013

José Fernando Seabra Babo

Screening molecular de Hepatozoon em anfíbios. Molecular screening of Hepatozoon in amphibian hosts.

Dissertação apresentada à Universidade de Aveiro para

cumprimento dos requisitos necessários à obtenção do grau de

Mestre em Biologia Aplicada, realizada sob a orientação científica

do Doutor D. James Harris, investigador do CIBIO-UP (Centro de

Investigação em Biodiversidade e Recursos Genéticos da

Universidade do Porto) e do Departamento de Biologia da

Faculdade de Ciências da Universidade do Porto e co-orientação

do Professor Doutor Amadeu Mortágua Velho da Maia Soares,

Professor Catedrático do Departamento de Biologia da

Universidade de Aveiro.

Universidade de Aveiro Departamento de Biologia

2013

José Fernando

Seabra Babo

Screening molecular de Hepatozoon em anfíbios. Molecular screening of Hepatozoon in amphibian hosts.

DECLARAÇÃO

Declaro que este relatório é integralmente da minha autoria, estando devidamente

referenciadas as fontes e obras consultadas, bem como identificadas de modo claro as

citações dessas obras. Não contém, por isso, qualquer tipo de plágio quer de textos

publicados, qualquer que seja o meio dessa publicação, incluindo meios eletrónicos, quer

de trabalhos académicos.

‘Não são as espécies mais fortes que sobrevivem

nem as mais inteligentes, e sim as mais suscetíveis

a mudanças.’

Charles Darwin

o júri

presidente Professor Doutor António José Arsénia Nogueira professor associado C/ Agregação da Universidade de Aveiro

Doutora Maria João Veloso da Costa Ramos Pereira bolseira de Pós-Doutoramento da Universidade de Aveiro

Doutor David James Harris investigador do CIBIO- Centro de Investigação em Biodiversidade e Genética

agradecimentos

Ao meu orientador James, por todo o tempo despendido comigo e pela ajuda

que me deu na realização deste trabalho. Foi um prazer trabalhar e apreender

consigo.

Ao Professor Amadeu Soares pela oportunidade que me deu em ir para o CIBIO

realizar a tese.

Ao João Maia por me ter ajudado nos primeiros passos deste trabalho e estar

sempre presente para ajudar, e por todo o conhecimento que me transmitiu.

A todas as pessoas do laboratório que tornavam o dia mais divertido e um ótimo

local de trabalho.

Um agradecimento especial a um amigo, Nuno Taser que me acompanha à sete

anos.

Aos meus amigos, Américo, André, Mimi, Neves, Zorion e muitos outros pelos

momentos de diversão passados durante o ano, e por me deixarem dormir na

sua casa quando precisava.

À minha namorada me conheceu na parte final deste projeto e me aturou e

apoiou sempre.

À minha tia Bina por todo o apoio que me deu.

À minha irmã pela ajuda com os esquemas e imagens, o lado artístico da família

ficou claramente com ela.

À minha mãe e a minha avó por todos os sacrifícios que fazem e fizeram para

que eu pudesse estudar e tirar um curso superior. Sem elas não era quem sou

hoje.

palavras-chave

resumo

Apicomplexa; parasita; Hepatozoon; anfíbio; Ichthyopthirius multifiliis; Dactylosoma ranarum; screening molecular; PCR.

A diversidade de parasitas tem sido pouco estudada devido a vários fatores, como o tamanho de algumas espécies, a localização dentro do hospedeiro, e o foco em grupos com fortes interesses antropogénicos ou com impacto econômico significativo. Apicomplexa é um grupo grande e diverso de parasitas unicelulares e com uma ampla distribuição e é o agente patogénico com mais sucesso conhecido pelo homem. Dentro do grupo alguns gêneros têm recebido pouca atenção, como caso de Hepatozoon. Estes são organismos pouco estudados que requerem mais amostragem. Técnicas de rastreio molecular, como PCR, são rápidas e fáceis de usar, e permitem a deteção eficiente desses parasitas. Elas também permitem a sequenciação produzindo dados para análises filogenéticas. Contudo mais dados são necessários para determinar corretamente a taxonomia e as relações filogenéticas do grupo. O objetivo deste trabalho foi detetar a diversidade de parasitas através de screening de amostras de tecido e sangue de diferentes espécies de anfíbios, utilizando primers específicos e reconstruir as relações filogenéticas das sequências produzidas. Várias espécies de anfíbios da Península Ibérica, Ilhas do Mediterrâneo, região Macaronésia e Marrocos foram analisadas usando primers específicos para parasitas e esfregaços de sangue foram observados. Parasitas foram encontrados em vários slides, sendo possível a identificação de pelo menos dois Hepatozoon, no entanto os primers não foram capazes de detetar as infeções. Apenas dois parasitas foram amplificados atráves de primers Hep, um parasita protozoário, Ichthyopthirius multifiliis, identificado numa amostra de Bufo calamita de Portugal, e um parasita Apicomplexa, Dactylosoma ranarum, em Pelophylax saharicus de Marrocos. Os primers utilizados não parecem ser úteis com anfíbios e novos primers precisam ser desenvolvidos para corretamente identificar a identidade dos parasitas observados nos slides.

keywords

abstract

Apicomplexa; parasite; Hepatozoon; amphibian; Ichthyopthirius multifiliis; Dactylosoma ranarum; molecular screening; PCR. Parasite diversity has been poorly studied due to several factors, like the size of some species, location within the host, and the focus on groups with strong anthropogenic interests or with significant economic impact. Apicomplexa is a vast and diverse group of unicellular parasites with wide distribution and the most successful pathogen known to man. Within the group some genus have received little attention, like the case of Hepatozoon. They are poorly studied organisms requiring more sampling. Molecular screening techniques like PCR are quick and easy to use, and allow efficient detection of these parasites. It also allows sequencing which produce data for phylogenetic analyses. Nonetheless more data is necessary to correctly establish taxonomy and phylogenetic relations of the group. The aim of this work was to assess the diversity of parasites trough screening of tissue and blood samples from different amphibian species using specific parasite primers and reconstruct the phylogenetic relationships of the sequences produced. Several species of amphibians from Iberian Peninsula, Mediterranean Islands, Macaronesia region and Morocco were analyzed using specific parasite primers and blood smears were observed. Parasites were observed in several slides, with at least two Hepatozoon being identified, however primers fail to detect them. Only two parasites were amplified, a protozoan parasite, Ichthyopthirius multifiliis, identified in a Bufo calamita sample from Portugal, and a Apicomplexa parasite, Dactylosoma ranarum, in Pelophylax saharicus from Morocco.

The primers used seem not to be useful with amphibians and new primers need

to be developed to correctly assess the identity of the parasites observed In the smears.

i

Index

Index…………………………………………………………………………………..I

List of Figures………………………………………………………………………...II

List of Tables………………………………………………………………………...IV

1 Chapter one - Introduction………………………………………………………….5

1.1 Parasitism…………………………………………………………………….6

1.2 Apicomplexa………………………………………………………………….7

1.3 Amphibian hosts……………………………………………………………19

1.4 Parasite detection techniques…………………………………………….23

1.5 Objectives…………………………………………………………………...29

1.6 Organization of the thesis………………………………………………….30

1.7 Taxonomy…………………………………………………………………...31

2 Chapter two - Materials and Methods……………………………………..……32

2.1 Sample collection…………………………………………………………..33

3 Chapter three - Putative Ichthyophthirius identified in the amphibian Bufo

calamita through molecular screening.…………………………………….……………….40

4 Chapter four - Screening for Apicomplexan parasites in amphibians…...…...48

5 Chapter five – General discussion………….……………………………………53

6 Chapter six - Concluding remarks and future possibilities…………………….57

6.1 Concluding remarks………………………………………………………..58

6.2 Future possibilities………………………………………………………….60

7 References………………………………………………………………………….61

Appendix 1…………………………………………………………………………….71

Appendix 2…………………………………………………………………………….72

ii

List of Figures

Figure 1.1 – Ultrastructural characteristics of the Apicomplexa. The image at the

bottom corresponds to an electron microscopy photograph of a Toxoplasma gondii. C -

conoid; CC - cortical cisternal layer; DG - dense granule; G – Golgi Complex; M -

microneme; N - nucleus; R - rhoptry. The parasite and vacuolar network (VN) are

enclosed by a vacuole membrane (VM). Scale bar, 1 μm. From:Kaasch and Joiner

(2000). The image at the top is a schematic of the principal cellular components of

apicomplexans. From: Šlapeta and Morin-Adeline

(2011)…………………………………………………………………………………………..10

Figure 1.2 – Life cycle of Apicomplexan parasites. The Center circle represents a

generic Apicomplexan life cycle. The outer circle represent the specific life cycle of

Plasmodium falciparum. The middle circle represents the life cycle of Toxoplasma

gondii. The bradyzoite form (∗) is responsible for reactivation of latent infection and is

an obligatory stage between tachyzoites and gametes. From: Morrissette and Sibley

(2002)…………………………………………………………………………………………11

Figure 1.3 – Hypothetical tree of Apicomplexa groups and their relationships. The width

and number on the branches refers to the named species and thus, the known

diversity. From: Šlapeta and Morin-Adeline (2011)…………….....................................13

Figure 1.4 – Bayesian tree representing Hepatozoon phylogenetic relations. Based on

562 bp 18S rRNA gene sequences. Bayesian posterior probabilities and ML bootstrap

values are given above and below the nodes respectively. + is indicated when both

values are 100%. The branches of JN181157, AF130361 and AF297085 were

shortened by 50%. From: Maia et al. (2012b)……………………………………………..18

Figure 1.5 – Diagram of the cycle of Hepatozoon sipedon. 1 Gamonts in erythrocytes of

the snake host are ingested by mosquitoes and are released in the gut. 2

Microgamonts and macrogamonts associate in syzygy in a parasitophorous vacuole in

a fat body cell of the mosquito haemocoel. 3 Gamonts undergo gametogenesis by 4

days post-feeding, after which one of the microgametes fertïlizes the macrogamete. 4

Resulting zygote forms an immature oocyst. 5 Nucleus of oocyst divides during the

initial stages of sporoblast development at 20 days post-feeding. 6 Oocyst, mature at

28 days post-feeding, contains an average of 600 sporocysts. 7 Each sporocyst

contains eight sporozoites. 8 Sporozoites are released into the gut of a frog when an

infected mosquito is ingested. 9 Dizoic cysts form in frog hepatocytes at 7 days post-

infection. 10 Cystozoites are released into the gut of a snake when an infected frog is

ingested. 11 Mature macromeronts are present in snake hepatocytes and other cells of

visceral organs after 15 days post-feeding. 12 Macromerozoites released from these

iii

macromeronts invade the bloodstream of the snake and reinfect hepatocytes and other

cells of visceral organs at 16 days post-feeding. 13 Micromeronts are mature after 30

days post-feeding. 14 Micromerozoites released from micromeronts infect erythrocytes

of the snake host, forming gamonts which are infective to mosquitoes during

subsequent feedings. From: Smith (1998)…………………………………………………22

Figure 1.6 – Representation of the life cycle of Hepatozoon catesbianae in his hosts. A.

Merozoites released from hepatic meronts enter erythrocytes. B. Merozoites transform

into gamonts. C. Mosquitoes feeding on infected frogs ingest erythrocytic gamonts. D.

Gamonts escape from erythrocytes in gut of mosquito and enter Malpighian tubules. E.

Micro- and macrogamonts come to lie within a common parasitophorous vacuole in

tubule cells. F. Gametogenesis ensues with formation of two biflagellate microgametes,

one of which fertilizes the macrogamete. G. The zygote expands into a spherical

oocyst. H. Oocysts undergo segmentation to form sporoblasts. I. Sporoblasts transform

into sporocysts. J. Each sporocyst contains four sporozoites. K. Frogs are infected by

ingesting mosquitoes containing sporocysts. L. Sporozoites enter hepatic parenchymal

cells where they develop into meronts. From: Desser et al. (1995)……………………..23

Figure 1.7 – Diagram representing the organization of nuclear genes of ribosomal

subunits. Scheme by José Babo……………………………………………………………29

Figure 2.1 – Geographic distribution of the species used as samples………………….33

Figure 2.2 –Photos of the species used during this thesis. A –Bufo bufo; B –

Amietophrynus mauritanicus; C –Discoglossus sardus; D –Bufo calamita; E –Hyla

meridionalis; F –Hyla sarda; G –Pelobates cultripes; H –Pelophylax perezi; I –

Pelophylax saharicus; J –Pleurodeles waltl; K –Bufotes balearicus; L –Bufotes

boulengerie. Photos E and I were taken by Daniele Salvi, Photos A, C, D, F, G, H, J, K,

L were taken by Matt Wilson, Photo B was taken by Pierre-Yves Vaucher……………35

Figure 3.1 – ML tree of the apparent Ichthyophthirius from a Bufo calamita, and closest

available comparative sequences from GenBank. Support for the Bayesian and for ML

analysis are given above and below the nodes, respectively. The branch of

Paramecium tetrawelia was shortened 75%.................................................................43

Figure 2.1 –Images of positive Hepatozoon infections in A) A. mauritanicus, and B) P.

saharicus, both of which failed to amplify using the screening protocol employed, C) a

positive infection of presumed Dactylosoma ranarum in Pelophylax saharicus, and D) a

typical negative sample. The scale bar corresponds to 20μm…………………………...50

iv

List of Tables

Table 2.1 –List of amphibians host species used in this thesis…..……………………..34

Table 2.2 –Details of the PCR primers used in this thesis……………………………….36

Table 2.3 –Reagents used, and respective concentrations for the Hep, HEMO and CR

primers, using invitrogen Taq DNA Polymerase…………………………………………..36

Table 2.4 – PCR protocols for the Hep, HEMO and CR primers using invitrogen Taq

DNA Polymerase……………………………………………………………………………...37

Table 2.5 – Reagents used, and respective concentrations for the Hep, HEMO and CR

primers, using Bioline MyTaqTM DNA Polymerase………………………………………..38

Table 2.6 – PCR protocols for the Hep, HEMO and CR primers using Bioline MyTaqTM

DNA Polymerase……………………………………………………………………………...38

Table 4.1 –Amphibian hosts species screened for parasites, the number tested using

alternative source material (tissue or blood), and the number examined under the

microscope on slides…………………………………………………………………………48

5

1 Chapter one

General Introduction, Objectives

6

1.1 Parasitism

The relations established between species are normally defined by the effect of the

interaction on each of the species. This relation can be beneficial for both species

(Mutualism), beneficial for one of the species but with no harm to the other

(Commensalism), beneficial for one of the species at the expense of the other (Parasitism,

Predation), one of the species can be inhibited with no effect on the other (Amensalism),

and finally both species can be inhibited (Competitive interactions).

In this thesis we are going to focus our attention on the relations of parasitism.

Therefore it is important to understand what a parasitic relationship means. If we go to the

origin of the word parasite, it derives from the Medieval French word parasite, which

comes from the Latin parasites, the latinisation of the Greek παράσιτος (parasitos), παρά

(para), "beside, by"+ σῖτος (sitos), "wheat". The word παράσιτος means “one who eats at

the table of another”. The literal interpretation leads to the conclusion that a large part of

living creatures are parasites (Poulin & Morand 2000). Bringing these to the context of this

thesis we could interpret parasitism as one living creature that feeds from another living

one without causing death to the latter, or else it would be predation.

Even though defining parasitism can be difficult and can lead to several different

classifications such as organism that lives in or on another living organism obtaining from

it part or all of its organic nutrient, and commonly exhibiting some degree of adaptive

structural modification (Bush et al. 2001). Therefore more strict definitions should be

adopted such as, a close host-parasite relation with the latter passing a large part of its life

history in or on his host. Even with the adoption of stricter definitions, the number and

diversity of parasites is huge (Poulin & Morand 2000). We can consider parasites within

taxa including bacteria, virus, fungi, algae, metazoans.

After classifying what parasitism is another problem emerges: How to classify the

types of parasites? This can be a hard task, and again several answers could be given.

We can divide them accordingly to where they can be found, endoparasites and

ectoparasites. Therefore the ones living inside the host body are endoparasites, and the

ones who live outside are ectoparasites. We can also classify them according to their size,

microparasites and macroparasites. Microparasites as the name says are microscopic,

while the macroparasites are bigger and can be seen without using the microscope. Both

micro and macroparasites can be either endo or ectoparasites. In this thesis we aim to

concentrate in a group of parasites that lives in the blood cells of their host, and therefore

are endoparasites and microparasites.

7

The size of parasites is one of many reasons why describing their diversity is so

difficult; some species can have very small sizes. Another reason is the primarily focus on

species considered valuable to man such as those with agricultural, veterinary or medical

interest (Poulin & Morand 2000; Morrison 2009). Some parasites occur with low

prevalence values, so inadequate sample efforts can also lead to an underestimation of

diversity values (Poulin & Morand 2000). Also, parasite identification has been traditionally

done with the use of the microscope. However, when identification is solely based on

morphological characteristics, this can lead to reports of distinct parasite species as a

single one, and vice-versa. In other words, many different parasites may morphological

appear very similar and be mistaken for a single species, known as “cryptic species”.

Even though this problem is starting to be overcome through the use of molecular

techniques, we can expect that the number of parasitic species described only represents

a fraction of the real diversity of this group (Adl et al. 2007; Morrison 2009). Another

problem is we still have not identified all free-living organisms in the world, and

considering that each of these organisms are potential hosts for at least one species of

parasite, we can expect that the diversity of parasites to be much higher than is currently

recognized (Poulin & Morand 2000).

1.2 Apicomplexa

The phylum Apicomplexa, also known as Sporozoa, comprehends a huge and

diverse group of unicellular protozoans with a wide environmental distribution. Most of

them are obligate intracellular parasites, and probably the most successful pathogens

known to man (Sato 2011). The morphological shape depends on the genus and lifecycle

stage and these parasites are typically quite host specific. They are known to parasitize a

large number of organisms, virtually all vertebrates, including humans, and marine and

terrestrial invertebrates (Frölich et al. 2012). Six thousand species are described, which

only represent a tiny fraction of the real number existing, estimated at 1.2 to 10 million

(Adl et al. 2007). This number can be very inaccurate, since it is believed that all animal

species host at least one of these parasites (Morrison & Ellis 1997). Not all biodiversity

has been described, and therefore with each new host identified, a potential new parasite

can be discovered (Poulin & Morand 2000).

The group can be found in humans and domestic animals, and is responsible for

several diseases with both medical and veterinary significance (Massimine et al. 2005).

Even though many of them are not pathogenic to their host, it is estimated that they cause

8

the deaths of 1 million people every year and agricultural losses of over US $1 billion per

year (Beck et al. 2009).

One of the most notorious parasitizing humans is the genus Plasmodium, the

agent responsible for malaria. It is estimated that the genus include about 172 species,

with 89 occurring in reptiles, 32 in birds and 51 in mammals, of which 4 cause malaria in

humans (Paul et al. 2003). The benign tertian malaria caused by Plasmodium vivax, is the

most widely distributed human malaria, with an estimated 70-80 million cases per year

(Cui et al. 2005), while Plasmodium falciparum is responsible for 3 to 5% of deaths

worldwide each year (Black et al. 2005), and is the most studied species in the genus.

Another important pathogenic to man is Toxoplasma gondii, an opportunistic parasite

present in 30% of the humans. The parasite represents little or no harm to healthy

individuals, but can be fatal in those with compromised immune systems, like AIDS or

cancer patients, and is dangerous for pregnant women (Hill & Dubey 2002; Massimine et

al. 2005). This pathogen is widely distributed across the globe, although its prevalence

varies with region (Hill & Dubey 2002). Cryptosporidium are widespread intestinal

pathogens and cause a disease called cryptosporidiosis (Beck et al. 2009). This disease

results in sickness and severe diarrhea, and in risk groups, like young children, the elderly

and immunosuppressed individuals, the disease can be fatal (de la Parte-Pérez et al.

2005). Other species infecting humans include Babesia, Cyclospora and Sarcocystis

(Leander 2003).

Apicomplexan parasites also cause huge agricultural losses, with Eimeria spp.

being responsible for losses over US $1.5 billion losses (Sharman et al. 2010). Eimeria

spp., which causes the disease coccidiosis has a huge economical importance for the

poultry industry (Beck et al. 2009; Frölich et al. 2012). Also Theileria, a tick-transmitted

Apicomplexan parasite, is known to infect livestock and causes important economic

losses (Beck et al. 2009). Neospora caninum is another pathogen of animals, responsible

for important losses in cattle. As the name suggest, the parasite was primarily associated

with dogs. Its cyst-forming parasite causing neuromuscular disorders in dogs and, a huge

cause of abortion and neonatal mortality in cattle (Dubey et al. 2007). Other relevant

parasites include Babesia, Besnoitia, Cryptosporidium, Sarcocystis, and Toxoplasma

(Muller & Hemphill 2013).

These parasites present very different transmission modes. Plasmodium and

Theileria for instance are vector-borne; Eimeria, Toxoplasma and Cryptosporidium form

highly resistant cyst that can be transmitted through contaminated materials, like food or

water (de la Parte-Pérez et al. 2005; Beck et al. 2009). Allied to this, the resistance to

9

most known drugs and the small number of existent vaccines makes very difficult to

prevent these diseases (Frölich et al. 2012).

Even with this important medical, veterinary and economical component many

other groups along with the Apicomplexa are very poorly studied. Although some

characteristics are easily studied such as life-cycles patterns, cyst organization, and

ultrastructure and host, the difficulty in finding and identifying these unicellular

endoparasites complicates the description of them (Morrison 2009).

1.2.1 General biology and life cycle

The name of the group Apicomplexa was determined by unique internal structures

that the organisms of the group possess (Levine 1973) (Figure 1.1). These structures

were only possible to observe after the invention of Electron Microscopy. The name of the

group derives from the presence of an apical complex on at least one of the life stage (Adl

et al. 2005). This complex is found normally in the infective stages at the front end,

displacing the nucleus and mitochondria towards the back (Aikawa et al. 1978). It is

responsible for recognizing, attaching and invading host cells (Smith & Desser 1997;

Walker et al. 2011; Frölich et al. 2012). The complex consists of a cytoskeleton

comprising a closed conoid and at least one polar ring, associated with secretory

organelles, rhoptries and micronemes (Adl et al. 2005; Walker et al. 2011). It also contains

Apicoplasts, a non-photosynthetic relict plastid (Walker et al. 2011), dense granules on

the posterior part, an endosymbiotic derived organelle mitochondrion, and the

acidocalcinomes. Apicomplexans also possess tubular mitochondrial cristae, micropores

and a pellicle with three membranous layers subtended by microtubules, which place

them within the Alveolates (Smith & Desser 1997). The number of and shape of rhoptries,

micronemes and dense granules vary according to the group (Leander 2008). Further, the

conoid is not present in the hematozoan group.

The life cycle is complex and can vary within the group (Morrissette & Sibley

2002). Sexual and asexual reproduction is present and several hosts can be involved

during this process. Having infected the host, the parasites invade the cells and divide

until the host cell is lysed and new parasites are released. Extracellular division normally

does not occur, therefore when released, these parasites need to invade new cells in

order for the cycle to continue (Morrissette & Sibley 2002).

10

Fig

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11

The life history normally consists at three distinct steps of development:

gametogony (sexual), sporogony (asexual) and merogony (asexual) (Leander 2003). A

sexual reproduction phase occurs, gametogony, where the fusion of gametes originates a

diploid zygote. This zygote invades a cell and rapidly originates haploid offspring through

meiosis. This process is called sporogony. After that, the haploid progeny, sporozoites,

leave the host cell to invade others specific cells. They use a variety of molecular tools,

including surface adhesions such as parasite surface proteins and binding antigens to

enter host cells (Baum et al. 2008). Upon establishment in a new host cell, sporozoites

typically produce merozoites, although pathway may differ depending on the species

(Figure 1.2). During this process, merogony, merozoites multiply, invading new cells and

giving origin to new merozoites. These merozoites can develop into gametocytes,

producing gametes so the cycle may continue. A scheme of a generic, Toxoplasma, and

Plasmodium life cycle is presented in Figure 1.2. Some aspects of the life cycle may be

different depending on the species (Adl et al. 2005; Frölich et al. 2012).

Figure 1.2 – Life cycle of Apicomplexan parasites. The Center circle represents a generic Apicomplexan life cycle. The outer circle represent the specific life cycle of Plasmodium falciparum. The middle circle represents the life cycle of Toxoplasma gondii. The bradyzoite form (∗) is responsible for reactivation of latent infection and is an obligatory stage between tachyzoites and gametes. From: Morrissette and Sibley (2002).

12

1.2.2 Phylogeny

The phylogeny of the Apicomplexa, and the groups within, is a controversial

subject, since in recent years establishes of relationships have been altered and

redefined. This chapter will present the current knowledge about Apicomplexa phylogeny,

followed by the phylogeny within the phylum.

Apicomplexa belongs to the monophyletic group Alveolata, along with

Dinoflagellates, Ciliates, and some minor lineages (Fast et al. 2002; Leander & Keeling

2004). Within the Alveolata, Apicomplexa and Dinoflagellates are more closely related to

one another than either is to Ciliates. Despite all the morphological differences between

Apicomplexa and Dinoflagellates, molecular tools support this phylogeny, and together

they form the Myzozoa (Escalante & Ayala 1995; Fast et al. 2002; Leander & Keeling

2004). The large number of morphological differences could be explained by the lack of

morphological information of intermediate lineages that are now being detected by

molecular tools (Leander 2003). Molecular phylogenetic analyses of several protein genes

have shown that the closest sister lineage of Dinoflagellates are the Perkinsids, mollusks

and microeukaryotes parasites (Bushek et al. 2002; Saldarriaga et al. 2003; Leander &

Keeling 2004). Perkinsids are not specifically related to Colpodellids and Chromerids, in

fact phylogenetic analyses of small subunit rRNA sequences, have shown that these two

groups are sister groups to Apicomplexa (Kuvardina et al. 2002; Leander 2003; Moore et

al. 2008). Colpodellids were suggested to be the earliest divergent sister group to the

Apicomplexa (Leander et al. 2003). The same conclusion was reached by Siddall et al.

(2001), using SSU rDNA to perform a phylogenetic analysis. Colpodellids are small

predatory flagellates that possess an apical complex used to consume algae and other

protists. Although the group presents an apical complex, it lacks the parasitic life style

typical of the Apicomplexa. This brings controversy into the phylogeny of the group, with

some authors considering that they belong with the Apicomplexa, while others consider

them a separate group (Adl et al. 2005; Walker et al. 2011). The phylum Chromerida was

also reported as a sister group of the Apicomplexa (Moore et al. 2008). This was

supported by the analyses of nuclear LSU rDNA and SSU rDNA sequences, and analyses

of the plastid rDNA (Moore et al. 2008). Despite being a photosynthetic alveolate, like

many Dinoflagellates, the photosynthetic plastid of this group is related most closely to the

apicoplast of Apicomplexa (Moore et al. 2008). Therefore Apicomplexa are currently

thought to be closer related to Chromerids than to Dinoflagellates, and this supports the

idea that the apicoplast is a trace of what remains of a red-algal derived chloroplast

(Waller & McFadden 2005; Janouškovec et al. 2010; Sato 2011).

13

The phylogeny within the Apicomplexa group was primarily estimated using

morphological characteristics. The use of molecular tools to support morphological traits

is in its infancy (Leander 2003). It has been postulated that before using any

morphological trait to establish taxonomy, the association with that group must be first

confirmed with clade analyses (Morrison 2009). The lack of support for morphological

traits used in the past, in conjunction with inappropriate taxon sampling and misuse of

genetic analysis tools are some reasons for earlier conflicting estimates of the phylogeny

of Apicomplexa (Kopecna et al. 2006; Morrison 2009). However, an attempt to improve

classification criteria is ongoing (Adl et al. 2007; Imam 2009). This could lead to changes

in the historically recognized groups, Coccidian, Cryptosporidia, Gregarines,

Haemosporinids, and Piroplasms (Barta et al. 2012). Figure 1.3 demonstrates a

hypothetical tree of the Apicomplexa.

Coccidians in conjunction with Haemosporinids and Piroplams form a clade. The

Coccidia group is a very diverse one, with many life cycles presented. Barta et al. (2012)

identify the tissue coccidia (Eimeriorina: Sarcocystidae), the enteric coccidia (Eimeriorina:

Eimeriidae), the adeleorinid coccidia (Adeleorina: Adeleidae), and the hemogregarines

(Adeleorina: various families). They can present monoxenous life cycle parasitizing

Figure 1.3 – Hypothetical tree of Apicomplexa groups and their relationships. The width and number on the branches refers to the named species and thus, the known diversity. From: Šlapeta and Morin-Adeline (2011).

14

vertebrate and invertebrate hosts (Eimeriidae and adeleorinid coccidian respectively).

They can also have heteroxenous life cycle (hemogregarines and Sarcocystidae), using

vertebrates as intermediate and invertebrates as definitive hosts. The mode of

transmission between hosts is usually a predator-prey relationship, infective stages are

produced within the prey and life cycle only completes within the predator. Like many

other Apicomplexa groups phylogeny may be poorly estimated, due to lack of enough

data and limited sample, with many organisms being overlooked, in favour of veterinary

important species. The remaining two groups, Haemosporinids and Piroplams are

considered sister clades and together form a monophyletic class called hematozoa (syn.

Aconoidasida) (Escalante & Ayala 1995; Adl et al. 2005). Both possess heteroxenous life

cycles, parasitizing vertebrate (sexual reproduction stages) and invertebrate (asexual

reproduction stages) hosts. The order Piroplasmida contains the genera Babesia and

Theileria, while the order Haemospororida contains the most medical and veterinary

significant genera Plasmodium, Haemoproteus and Leucocytozoon. An endosymbiotic

marine protist with uncertain classification since its discovery in the 19th century is the

newest addition to the Apicomplexa and Hematozoa: Nephromyces are ubiquitous

nonhereditary symbionts, transmitted horizontally to new hosts. This relation has been

established using rDNA and morphological traits (Saffo et al. 2010). Cryptosporidia are

intracellular monoxenous parasites that infect vertebrates, including humans. They have a

direct life cycle, with intracellular but extracytoplasmatic development. The morphology

and life cycle are typically Coccidian, and Cryptosporidium was considered a member of

coccidia, until phylogenetic evidence showed its closer affinity with gregarines (Zhu et al.

2000; Leander et al. 2003; Leander & Keeling 2004). The group possibly evolved from

Gregarines, however the position within the Apicomplexa remains uncertain (Rueckert et

al. 2011). Some fundamental differences to Coccidians are evident: the lack of a plastid;

the presence of an acristate, ribosome-studded mitochondrion posterior to the nucleus;

and plant-like polyamine biosynthesis by decarboxylation of arginine rather than ornithine.

Another big difference is the resistance to anticoccidial drugs, which would not be

expected if Cryptosporidia belonged with Coccidia. The distinction of the two groups

explains why this resistance was observed (Zhu et al. 2000). The last group is the

Gregarines, which inhabit different body spaces within the marine and terrestrial

invertebrate hosts. They possess monoxenous life cycle and can present extracellular

feeding stages. Gregarines are considered to be product of the first lineage that diverges

within the Apicomplexa. It is the less well known group within the Apicomplexa. The lack

of information on this group makes unclear whether Cryptosporidium should be nested

15

within Gregarines, although the majority of phylogenetic studies are based on the small-

subunit ribosomal DNA sequences from a few species (Leander 2003; Rueckert &

Leander 2008), which may explain this lack of resolution.

The phylogeny and taxonomy within this ancient lineage therefore still remains

uncertain. Apicomplexa probably diverge from Dinoflagellates 700-900 Million years ago

(Escalante & Ayala 1995; Douzery et al. 2004). Despite several revisions, controversy

remains within the group. The recent placement of Nephromyces reveals that not all

apicomplexans present parasitic life style. This show how much is still to be discovered

about this particularly diverse group of organisms.

1.2.3 Hepatozoon

Hepatozoon species (Apicomplexa: Adeleida) are hemogregarines with

heteroxenous life cycle known to parasitize most groups of tetrapod vertebrates as

intermediate hosts and a large number of blood sucking invertebrates (ticks, mites, lice,

fleas, reduviids and dipterans) as definitive hosts (Smith 1996; Harkness et al. 2009).

About 336 species of Hepatozoon have been reported (Smith 1996). They can be found in

the visceral organs and blood cells of the hosts (Criado-Fornelio et al. 2007). The way of

transmission is trough arthropods vectors (Smith 1996). Other routes are known to occur,

like vertical transmission in the case of H. canis (Murata et al. 1993) and prey predator

transmission when the intermediate vertebrate host ingests the definitive host containing

Hepatozoon oocysts. Species within the genus share some basic characteristics in their

life cycle such as: an asexual stage, sporogony, occurs in a haematophagous invertebrate

definitive host; the merogony and the sexual stage, gamontogony, occurs in the vertebrate

host (Baneth et al. 2003). There are some doubts about the taxonomic placement of the

genus Hepatozoon. Originally assigned within the family Haemogregarinidae (Lèger

1911), the genus was then elevated to family level by Wenyon (1926), just to be assigned

as a genus again by Levine (1988). To fully assess specific status, morphological

characters, life cycle patterns and host specificity is required (Mathew et al. 2000; Perkins

& Keller 2001). Nevertheless, much of this information is not collected, and lack of

information about the sporogonic development exist (Baneth et al. 2003; Moço et al.

2012). Thus, same species can be encountered with different names just because they

are found in another location or different host (Smith et al. 1999).

Both Haemogregarina spp., Hepatozoon spp., and Karyolysus spp., present

similarities in the intraerythrocytic gamonts, and thus were all placed within the family the

large family Haemogregarinidae (Baneth et al. 2003). However, differences in the vector

16

choice of these genus justify the separation into different families (Baneth et al. 2003).

Parasites transmitted by the bite of leeches and found in cells of cold-blooded vertebrates

represent the family Haemogregarinidae (Haemogregarina, Cyrilia and Desseria spp.). If

the vectors are ticks or mites and they can be found parasitizing cold blooded vertebrates

they should represent the family Karyolysidae (Karyolysus and Hemolivia spp.). If the host

in infected by resilient oocysts with numerous sporocysts by the ingestion of an

invertebrate host, characteristic of Hepatozoon spp., it should be placed within the family

Hepatozoidae. The hypothesis that the genus Hepatozoon should raised to a family level

was supported by Barta et al. (2012). In their study based on 18S rDNA, they suggest that

the genus should be raised to family level, or, as already proposed by Smith and Desser

(1997), using morphological characters, divided into at least two genera, making

Hepatozoon paraphyletic. The passage from a monophyletic to a paraphyletic group was

also supported by Mathew et al. (2000). In their study they report three different

associations, one where Hepatozoon aegypti and Hepatozoon gracilis were clustered

together, both species uses mosquitoes as vectors; the second one presents Hepatozoon

americanum and Hepatozoon canis, again the two parasites use the same type of vector,

ixodid ticks; the third one presents Hepatozoon lygosomarum in a clade together with

representatives of Haemogregarina, Cyrilia, Desseria, Karyolysus, and Hemolivia.

Nevertheless this was achieved using morphological characters, which are known to be

homoplastic. Barta et al. (2012) reported 4 different clades for Hepatozoon spp., including

one sequence of Hemolivia mariae. The clades present high degree of host-parasite

association of various species with their definitive hosts. The association with the

vertebrate host could be there could be lower. First the most basal clade includes species

of Hepatozoon canis, Hepatozoon americanum, Hepatozoon ursi, Hepatozoon felis, and

an unnamed Hepatozoon sp. from the pine marten. Thus these Hepatozoon species

typically use carnivores and ixodid ticks as hosts. The next diverged clade consisting of an

unnamed Hepatozoon species and the Hemolivia mariae. The host in this clade are

reptiles (brown water python and Australia sleepy lizard respectively), and Amblyomma

species (ticks). The third group presented comprised Hepatozoon species infecting

marsupial mammals, and using Ixodes species (ticks) as definitive host. The final clade is

the most derived one, it present Hepatozoon species using a variety of amphibians,

rodents and reptiles as intermediate host, and several arthropods as vectors. Similar

phylogenetic trees have been achieved in other studies (Figure 1.4) (Harris et al. 2011;

Pinto et al. 2012). However Harris et al. (2013a) results contrary the ones of Barta et al.

(2012), and suggest that Hemolivia should be not included with Hepatozoon. The small

17

size of the sequence used by Barta et al. (2012) could compromise estimates of

phylogenetic relationships. More details are necessary to correctly infer phylogeny within

Hepatozoon. Although parasites found in reptiles and snakes do not form a monophyletic

group, amphibians seems to (Figure 1.4). Within the hemogregarines, Hepatozoon

species possess the most complex life cycles. The heteroxenous life cycle probably

evolved from a monoxenous ancestral. However, it is not establish if they evolve to two

host life cycle and from that to three host, or the reverse occurred. In the final comments

of their work, Barta et al. (2012) states the need to gather more data about life cycle data

additional sequences from other species, to better understand their evolutionary history.

The first report of a bat infected with a Hepatozoon species was made by Pinto et

al. (2012). A large number of bats species have an insectivorous diet, so prey-predator

patterns are possible. However some species also predate on small vertebrates which

can potentiate the opportunity for transmission. The prey-predator transmission route has

been reported in several studies, especially in carnivores and snakes (Baneth et al. 2003;

Allen et al. 2011; Baneth 2011; Tomé et al. 2012; Viana et al. 2012). Baneth (2011)

suggests that Hepatozoon americanum infection in dogs may have two transmission

routes.

Hepatozoon americanum is responsible for American canine hepatozoonosis

disease, the parasite can be transmitted by the Gulf Coast Tick (Amblyomma maculatum)

or by predation and ingestion of parasite cystozoite forms from mammal host tissues.

Other transmissions pathways are possible in Hepatozoon species (Murata et al. 1993;

Baneth 2011; Hornok et al. 2013). Vertical transmission of H. canis has been proved,

Murata et al. (1993) kept naturally infected pregnant females in a free controlled

environment until the birth occur. They were able to observe meronts and gamonts in the

progeny few days after the birth.

Amphibians are part of many Hepatozoon species life cycles, either as unique

intermediate host, or as a first intermediate host later ingested by a second one (Smith et

al. 1994; Kim et al. 1998; Smith et al. 1999; Viana et al. 2012). The number of stages of

life cycle within the amphibian host may differ according to the Hepatozoon species. In a

three host pattern transmission, sporogony occurs in the haemocoel of the definitive host

(Smith 1996). After that, the first intermediate host (many times an amphibian) ingests a

mosquito and development of cysts occurs in the liver and lung tissues (Smith 1998;

Viana et al. 2012). The second intermediate host consumes the parasitized paratenic host

and cystozoites are released giving origin to meronts. Paratenic hosts are hosts that are

not necessary for the development of a particular species of parasite, however they could

18

Figure 1.4 – Bayesian tree representing Hepatozoon phylogenetic relations. Based on 562 bp 18S rRNA gene

sequences. Bayesian posterior probabilities and ML bootstrap values are given above and below the nodes respectively. + is indicated when both values are 100%. The branches of JN181157, AF130361 and AF297085 were shortened by 50%. From: Maia et al. (2012b).

19

be used to maintain the life cycle of that particular parasite. Merozoites are then released

into the blood stream and infect erythrocytes giving origin to gamonts. These gamonts are

ingested by the mosquito when he feeds from the blood of the second intermediate host.

This case is reported in caimans and snakes (Smith 1998; Sloboda et al. 2008; Viana et

al. 2012). The difference to two host life cycles is the absence of a cystic stage. The

sporocysts ingested by the amphibian release sporozoites that directly give origin to

meronts. Meronts give origin to merozoites and the process proceeds identical to the

previous described (Desser et al. 1995; Kim et al. 1998). Only one round of hepatic or

erythrocytic merogony is reported in frogs (Kim et al. 1998; Smith et al. 2000). Mature

gamonts resulting of invade erythrocytes by merozoites appear after some weeks time

(Harkness et al. 2009). Normally each erythrocyte only presents one gametocyte,

however more than one can also be found (Jovani et al. 2004).

The level of pathogenicity of Hepatozoon is still not clear. Mortality in mosquitoes

that feed on blood of infected hosts has been reported (Ball et al. 1967; Smith 1996;

Harkness et al. 2009). Three mosquito species (Culex. tarsalis, Anopheles albimanus, and

Aedes sierrensis) that fed on blood from indigo snakes (Drymarchon corais) infected with

Hepatozoon rarefaciens, presented considerable mortality (Ball et al. 1967). Two other

species, Culex territans and Culex pipiens, also presented the same result when fed from

garter snakes (Thamnophis sirtalis) infected with Hepatozoon sipedon (Smith 1996).

These results were reported with mosquitoes that fed on frogs as well (Harkness et al.

2009). However in the last case the cause of death was not confirmed. Information

regarding Hepatozoon species is still very poor is some areas, with most of the efforts

concentrated in medical and veterinary important species. There is a need to gather much

more data regarding life cycles, sequences from different species and genes, to be

possible to correctly establish taxonomy and phylogenetic relation of the group.

1.3 Amphibian hosts

Amphibians are the most threatened class of vertebrates worldwide (Koprivnikar et

al. 2012; Li et al. 2013). According to IUCN data nearly 41% of amphibian species are

threatened with extinction, that is classified in the Red List as ‘vulnerable’, ‘endangered’ or

‘critically endangered’ (Li et al. 2013). Habitat loss, pollution, invasive species, and various

pathogens such as Ranavirus, the chytrid fungus Batrachochytrium dendrobatidis, and

protistan parasites are some of the reasons for global extinctions and mass mortalities

(Aisien et al. 2011; Koprivnikar et al. 2012; Landsberg et al. 2013; Li et al. 2013).

20

Amphibians can be either definitive hosts or intermediate hosts for their parasites.

Definitive when they are the only vertebrate host used by the parasite in its life cycle,

intermediate when they are used as the first vertebrate host and are the vehicle of

transmission to the vertebrate that going to be the definitive host. Macroparasites and

microparasites require different resources from the host.

Many studies have been carried out on amphibian parasites. For example, a study

conducted with Leptodactylus melanonotus in México, reported 20 taxa of helminths

infecting this species (7digeneans and 13 nematodes). This raises the number of

helminths parasitizing L. melanonotus to 36 (Mata-López et al. 2012). Recently

trematodes parasites, known to play important ecological roles in the aquatic environment,

have been investigated as sources of pathology and mortality in amphibians (Orlofske et

al. 2013; Preston et al. 2013). The most commonly found echinostome trematode

Echinostoma trivolvis causes negative effects in growth and development, and depending

on the development stage and age of larval can be fatal (Szuroczki & Richardson 2012;

Orlofske et al. 2013). Another trematode parasite, Ribeiroia ondatrae, is responsible for

malformations such as missing limbs or extra limbs (Lunde & Johnson 2012). In Benin

several species of trematodes (five species) were also reported in amphibians, as well as

eight nematodes species, three monogeneans, and two cestodes (Aisien et al. 2011).

These parasites can be found in several different organs and body cavities. Hosts with

different ecology should present different parasites. Usually nematodes are found in

relatively terrestrial amphibian species, while on the other hand trematodes are normally

found in ranids, tree frogs, and aquatic amphibians (Koprivnikar et al. 2012).

Virus are also important pathogens for amphibians, and an important one

responsible for population decline is the Frog virus 3, a species of the genus Ranavirus,

that infects both larval and adult amphibians (Landsberg et al. 2013). Funguses are also a

parasite of amphibians. A pathogenic and perhaps the most famous one is the chytrid

fungus, Batrachochytrium dendrobatidism. It is responsible for a fatal and infectious

disease called chytridiomycosis (Li et al. 2013). The fungus infects the amphibians

causing electrolyte imbalance through disruption of cutaneous osmoregulatory functions

which can result in death (Li et al. 2013).

Occupying both an aquatic and terrestrial environment, amphibians are exposed to

a huge variety of vectors and consequently they are very susceptible to acquire blood

parasites (Barta & Desser 1984). Amphibians are known to accommodate a variety of

blood parasites such as Apicomplexans, filarial nematodes, hemoflagellates, bacteria, and

viruses (McKenzie & Starks 2008).

21

In Australia two Myxosporean Parasites, Cystodiscus axonis and Cystodiscus

australis, have been reported in seven Australian frogs species, of which four are

endangered species (Hartigan et al. 2012a; Hartigan et al. 2012b). Cystodiscus parasites

are associated with inflammation of the nervous tissue and hepatic disease.

Infections with different species of Trypanosoma, have been reported in several

species of amphibians around the world (Barta & Desser 1984; Žičkus 2002; Readel &

Goldberg 2009; Gupta et al. 2012). In Costa Rica four Trypanosoma parasites have been

identified in frogs, two at the species level (Trypansoma loricatum and Trypanosoma

chattoni) (McKenzie & Starks 2008). The same authors with frogs from Uganda identified

besides Trypanosoma sp., a Hepatzoon sp., and a microfilariae of undetermined

classification. A nematode microfilariae (Foleyellides striatus) was also identified in frogs

from Costa Rica, and in the same study two Apicomplexans were reported (Hepatozoon

sp. and Lankesterella sp.) (McKenzie & Starks 2008).

Several species of hemogregarinas have been discovered from the family

Bufonidae. In India, a report of a Hepatozoon sp. in blood from a Bufo melanostictus, lead

the authors to try to classify the parasite within the ones discovered in the Bufonidae

family. However the lack of similarity gave rise to a novel Hepatozoon species, H.

gangwarii n. sp. (Gupta et al. 2012).

Within tetrapods, Apicomplexa are very common blood parasites, and one genus

is often identified, Hepatozoon, a major part of this thesis. Hepatozoon are known to

parasitize blood cells of several organisms including amphibians. Within this genus only

42 species are described in anurans, and from those only 2 have complete life cycles

described (Boulianne et al. 2007).. Hepatozoon caimanis, a parasite of Caiman yacare

and Caiman latirostris, seems to have anurans as intermediate host in the Pantanal region

(Viana et al. 2012). This Hepatozoon uses anurans as paratenic hosts, and the

transmission occurs when the crocodilians predate and eat these amphibians. Similar

transmissions occur with other parasite species. Hepatozoon sipedon was reported to

infect the Northern leopard frog (Rana pipiens), which is used as an intermediate host in

the transmission route to the Northern water snake (Nerodia sipedon sipedon) (Smith et

al. 1994). This species, as many others that use amphibians as intermediate hosts, has

three hosts during its life cycle. Cystic development takes part in an anuran host and

merogonic development occurs in snakes (Figure 1.5).

From frogs of the genus Rana seventeen species of Hepatozoon have already

been described (Smith et al. 2000). Among them the two species that have fully described

life cycles, Hepatozoon catesbianae and Hepatozoon clamatae.These two species of

22

Hepatozoon have been reported in frogs of Nova Scotia and other locations (Boulianne et

al. 2007). These authors observe higher affinity of Hepatozoon clamatae for green frogs

than for bullfrogs. The opposite was observed with Hepatozoon catesbianae, were

bullfrogs showed a higher affinity. The life cycle of these two species is very similar both in

the mosquito vector and vertebrate host (Kim et al. 1998) In comparison with the life cycle

of Hepatozoon sipedon, Hepatozoon catesbianae only has two hosts during the life cycle

(Figure 1.6). This species has a direct life cycle without a cystic stage (Smith et al. 1999).

Figure 1.5 – Diagram of the life cycle of Hepatozoon sipedon. 1 Gamonts in erythrocytes of the snake host are ingested by mosquitoes and are released in the gut. 2 Microgamonts and macrogamonts associate in

syzygy in a parasitophorous vacuole in a fat body cell of the mosquito haemocoel. 3 Gamonts undergo gametogenesis by 4 days post-feeding, after which one of the microgametes fertïlizes the macrogamete. 4 Resulting zygote forms an immature oocyst. 5 Nucleus of oocyst divides during the initial stages of sporoblast development at 20 days post-feeding. 6 Oocyst, mature at 28 days post-feeding, contains an average of 600 sporocysts. 7 Each sporocyst contains eight sporozoites. 8 Sporozoites are released into the gut of a frog when an infected mosquito is ingested. 9 Dizoic cysts form in frog hepatocytes at 7 days post-infection. 10 Cystozoites are released into the gut of a snake when an infected frog is ingested. 11 Mature macromeronts are present in snake hepatocytes and other cells of visceral organs after 15 days post-feeding. 12 Macromerozoites released from these macromeronts invade the bloodstream of the snake and reinfect hepatocytes and other cells of visceral organs at 16 days post-feeding. 13 Micromeronts are mature after 30 days post-feeding. 14 Micromerozoites released from micromeronts infect erythrocytes of the snake host, forming gamonts which are infective to mosquitoes during subsequent feedings. From: Smith (1998).

23

1.4 Parasite detection techniques

Parasitologists search for the most accurate method for detecting and identifying

parasites never stops, and nowadays there are several different tools at their disposal to

do this work. We can divide these tools into classical diagnostics techniques and nucleic

acid-based diagnostics. In the classical diagnostics techniques we can place microscopy,

Figure 1.6 – Representation of the life cycle of Hepatozoon catesbianae in his hosts. A. Merozoites released from hepatic meronts enter erythtocytes. B. Merozoites transform into gamonts. C. Mosquitoes feeding on infected frogs ingest erythrocytic gamonts. D. Gamonts escape from erythrocytes in gut of mosquito and enter Malpighian tubules. E. Micro- and macrogamonts come to lie within a common parasitophorous vacuole in tubule cells. F. Gametogenesis ensues with formation of two biflagellate microgametes, one of which fertilizes the macrogamete. G. The zygote expands into a spherical oocyst. H. Oocysts undergo segmentation to form sporoblasts. I. Sporoblasts transform into sporocysts. J. Each sporocyst contains four sporozoites. K. Frogs are infected by ingesting mosquitoes containing sporocysts. L. Sporozoites enter hepatic parenchymal cells where they develop into meronts. From: Desser et al. (1995).

24

and serology-based assays (Immunodiagnosis – antibody detection, Antigen detection). In

the nucleic acid-based diagnostics are multilocus enzyme electrophoresis, southern blot

technique, PCR, and LAMP (loop mediated isothermal amplification).

Microscopy is a classical diagnostic technique of common use that was the only

tool available to parasitologists in the past that allowed the detection and characterization

of microparasites (Ndao 2009). The use of this tool was only possible due to the work of

the Dutch scientist Antony van Leeuwenhoek, that turned the microscope from a novelty

to a scientific tool (de Waal 2012). It allows to diagnose infection in various host samples

(Ndao 2009).

The microscope was such an important tool, that the data allowing the first

taxonomic assignments and phylogeny reconstruction for parasites were based on this

technique. With the evolution of this tool (e.g. Electron microscopy), the knowledge about

parasites has also evolved, and more accurate data has been produced. The discovery of

many intracellular structures, impossible to see with the optical microscope, allowed better

reconstruction of the phylogeny of many parasites and also a better taxonomic

arrangement. In the case of haemosporidians the microscope has been used as a tool to

describe several aspects such as life history strategies, vertebrate hosts, and aspects of

ecology for more than 100 years (Valkiūnas et al. 2008). One of the simplest applications

of this technique consists in the examination of smears in a slide. In the case of

Protozoans in the circulatory system (Hepatozoon, and many other Apicomplexans) blood

smears are prepared. The negative aspect of this method is that usually the preparation

and examination of the samples is time-consuming, labour intensive, and the correct

diagnosis is dependent of experienced and qualified staff (Ndao 2009; de Waal 2012).

Microscopy has several attractive aspects, offering advantages over other methods. With

microscopy it is possible to quantify the intensity of the infection, easily identify mix

infections (different types of parasites within the same host), differentiate between the

distinct developmental stages of the life cycle, and determine which tissue or cell the

parasite is occupying. Nonetheless, the technique has limitations that can lead to wrong

taxonomic placements, failure to detect infections, especially when infection levels are low

and the difficulties of using morphological characters to reconstruct phylogenies (Richard

et al. 2002; Morrison 2009).

In a study conducted by Richard et al. (2002), they compared the efficiency of PCR

and microscopy for detection of avian haemosporidians. Their results showed that PCR is

more accurate detecting the presence of these parasites; they also reported that for

screening a large number of samples, PCR was faster, cheaper, and more reliable than

25

microscopic screening. Other studies reported the same superiority of PCR tests over

microscopy (Jarvi et al. 2002; Durrant et al. 2006). These results were not achieved by

Valkiūnas et al. (2008), who state that PCR and microscopy methods underestimate

roughly the same amount of infections in haemosporidian parasites, except in the case of

low infection levels. They also argue that the results presented by microscopy can be

influenced by the protocol used.

The search for more accurate methods led parasitologists to the serology-based

assays, that are more sensitive than microscopy and can be used to indirectly detect

infections (Ndao 2009). This method can be used when biological samples or tissue

specimens are unavailable or the parasites occur at very low densities.

Serological techniques are divided into two groups, the antibody-detection assays

and the antigen-detection assays. Common tests used to detect the presence of

antibodies in response to a specific protozoan in the antibody-detection groups are: the

complement fixation test (CFT); the immunodiffusion (ID); the indirect haemagglutination

(IHA); the latex agglutination (LA); the indirect or direct immunofluorescent antibody test

(IFAT or DFAt); the radio-immunoassay (RAI); and the enzyme-linked immunosorbent

assay (ELISA). These tests have some negative aspects such as false positives that can

result from cross reactions between closely related parasites. Another cause of false

positives is the long persistence of the antibodies, even after elimination of the parasite.

This means that not all positive results are resulting from an infected host (de Waal 2012).

Possible solutions for this problem are the antigen-detection assays. These tests, instead

of detecting the host antibodies, aim to specifically detect the parasite antigens. However,

these tests also have negative aspects. There are no standardized protocols and reagents

for these tests, which result in variation in the results between laboratories. Furthermore,

cross reactions continue to be presented (de Waal 2012). Both groups of tests have been

improving and can be found commercially.

A study using three different methods, microscopy, PCR and a serology based

method (IFAT) was performed by Karagenc et al. (2006). They were trying to detect

Hepatozoon spp., and at the same time determine which method was the most efficient.

They were able to identify Hepatozoon canis and the method that was presented to be the

most efficient detecting the parasite was the serologic method (36.8%), followed by PCR

(25.8%) and finally microscopy (10.6%).

The nucleic acid-based diagnostics techniques, or also called molecular tools, are

compared to microscopy, much more recent. The use of these tolls has recently become

standard in field surveys of parasites detection (Beck et al. 2009). As any other technique,

26

they present positive aspects but also some negative ones. PCR is by far the most

common of these techniques, but other techniques like Multilocus enzyme

electrophoresis, Southern blot technique and LAMP are also used (Ndao 2009; de Waal

2012). These methods have improved sensitivity and specificity over the classical ones

(Ndao 2009).

Multilocus enzyme electrophoresis consists in characterizing organisms by the

relative mobilities under electrophoresis of a large number of intracellular enzymes. The

method has been used in some studies with Trypanosoma and Apicomplexa parasites

(Shirley 1975; Barnabé et al. 2000). The disuse of this method compared to PCR comes

from the many disadvantages it possess such as, failed identifications, the impossibility of

knowing the degree of relationship between different phenotypes, and because it is a time

consuming and expensive method (de Waal 2012).

Southern blot technique uses restrictions enzymes to digest DNA fragments.

These fragments, after being separated by electrophoresis, are transferred onto

membrane filters and hybridized with complementary labelled probes. The method has

already been applied to several protozoans. Nonetheless the design of proper probes

capable of hybridizing and digest the DNA fragments is a limitation to the technique (de

Waal 2012).

LAMP has also been used in protozoans with some good results (Karanis &

Ongerth 2009). The method consists of using six different primers, purposely designed to

recognize eight independent regions of the target gene. Amplification only occurs if all the

primers bind and form a product. To amplify the target DNA a robust polymerase (BST) is

used, followed by an autocycling strand displacement mechanism at a constant

temperature (60-65ºC). Contrary to PCR, there is no need to perform variations in

temperature and DNA extraction is not necessary (Karanis & Ongerth 2009; de Waal

2012).

In 1983 the classical method of PCR was developed. This method is still by far the

most commonly used. There are many reasons for the popularity of this technique; it is

simple to use, fast, does not require much time, is sensitive and cheaper than the

conventional methods (Richard et al. 2002; Su et al. 2010). This technique consists of

denaturation by heating of the double-stranded genomic DNA template. After that a

decrease in the temperature allows the set of primers to hybridize (anneal) to their

complementary sequences. The next step is the extension of the template DNA in both

directions from the primer sites by enzymatic catalysis with a thermostable DNA

polymerase (Taq) and results in double-stranded products. A third step of higher

27

temperature denatures the DNA, and the cycle is then repeated normally 30 to 40 times.

The presence or not of amplifications of the expected sizes is interpreted as an infected

sample or not, respectively. The original PCR may be modified to further increase

sensitivity and specificity, such as the nested PCR, RT-PCR (real-time PCR), and

multiplexed PCR (Beck et al. 2009; Ndao 2009; de Waal 2012). These modifications were

used to try to overcome some of the limitations of the classical PCR.

Microscopy is able to easily detect mixed infections, and quantify the intensity of a

given infection. This is not as easy in PCR, and is one of the limitations of the method.

Each set of primers (depending in their specificity) detects a set of parasites in the PCR

reaction. To overcome this limitation detecting mixed infection, multiplex PCR has been

used. Combined use of numerous specific primer sets into a single PCR assay allows the

detection of different parasites in the same reaction (Zarlenga & Higgins 2001). Real-time

PCR has been used to try to solve the second limitation of the method. This variant of

PCR, besides reducing the problems of cross-contamination, allows the quantification of

infection intensity (Gasser 2006). The method has been used with different parasites,

including Hepatozoon spp.(Criado-Fornelio et al. 2007). PCR being a sensitive technique

is susceptible to cross-contamination and can produce false positives. If the primers

chosen have low specificity or the infecting parasites were very closely related, the other

parasites may also be amplified, which may be wrongly interpreted. The specificity of the

reaction is very important to avoid amplifications of other organisms that can be in the

host. The specificity is controlled by many factors, such as the design of the primers,

buffer and cycling conditions, particularly primer annealing temperature. Therefore

choosing or designing appropriate primers is crucial. This choice will be influenced by the

questions to be addressed. Genes possess different evolution rates, and consequently the

targeted region must contain an adequate sequence variability to allow the identification to

the taxonomic level required. On the other hand, highly conserved primers may amplify

the host. Sequencing all PCR products helps reduce this problem.

The possibility of generating sequences data for phylogenetic and epidemiological

studies of parasites makes PCR a very attractive method (Valkiūnas et al. 2008). The

sequences produced are the reflection of the region targeted. The targeted regions for

Apicomplexan parasites can be several, including mitochondrial genomes, nuclear

genomes, and apicoplast genomes (Hikosaka et al. 2013). All suffer and accumulate

mutations over time, and the rate of these mutations varies according to the region

(Gasser 1999).

28

Mitochondrial DNA is commonly used to study population genetics,

phylogeography, speciation, systematic, and genetic variation in many species and

genera (Geller et al. 2013). The choice for these genes is due to particularly

characteristics that they possess, making them attractive to perform phylogenetic

analyses. These characteristics include high copy number, conserved sites, and lack of

recombination since they are normally haploid and only inherited from the mother (Gasser

1999). However, the mitochondrial genome of Apicomplexa parasites has some

characteristics that make it hard to work with. Although they are greatly reduce,

sometimes possessing only three protein-coding genes (Cytochrome c oxidase subunits I

and III, and cytochrome b), this group of parasites presents high variability in the genome

structure, and also presents NUMTs (transfer of homologous mtDNA to nuclear DNA),

and rRNA fragments (Hikosaka et al. 2010; Hikosaka et al. 2013). Apicoplast genomes

seem to present high variability in size and structure. Complete sequences for these

genomes are available from 9 Plasmodium species, Babesia bovis, Theileria parva,

Eimeria tenella and Toxoplasma gondii (Hikosaka et al. 2013). In the case of nuclear

genomes, the sequences available report to species with medical or veterinary

importance, with complete sequences available from seven Plasmodium species, two

Cryptosporidium species, Babesia bovis, Theileria parva, Theileria annulata, Eimeria

tenella and Toxoplasma gondii (Walker et al. 2011; Hikosaka et al. 2013). Even with the

increasing number of complete genomes the production of new nuclear markers has not

been easy. Ribosomal RNA genes display several attractive characteristics to be used in

molecular studies such as the high number of copies in the genome; they have conserved

and variable regions, great characteristics for primer design and a good source of

phylogenetic data respectively; and they possess a high number of transcripts in the cell

(Perkins et al. 2011). rRNA is a good provider of functional genetic markers, and these

genes, especially 18S rRNA (small subunit ribosomal RNA), are the most commonly used

for phylogenetic reconstructions of protists (Adl et al. 2007). These genes consist of

tandemly arrayed sequence repeats, normally encountered in clusters, that can be found

in across the genome (Gasser 1999). The 18S rRNA is consider a good marker to be

used for reconstructing phylogenetic relationships among apicomplexans, and also

protists in general (Adl et al. 2007; Perkins et al. 2011). Nevertheless, Perkins and Keller

(2001) alerts to two possible setbacks: firstly 18S rRNA genes of Apicomplexa parasites

possess several insertions/deletions that can lead to difficulties in ascertaining alignment;

secondly several species of Apicomplexa parasites possess specific arrangements of

these genes, they can present variable number of copies dispersed among many

29

chromosomes, that are expressed at different stages of the life cycle (Perkins & Keller

2001). Furthermore, this gene is generally quite slowly evolving and so may not be

informative at distinguishing between more closely related parasites.

Other markers have been applied to parasites studies. Internal transcribed spacers

(ITS) offer accurate species markers (Gasser 1999). These genes (ITS1 and ITS2) are

highly variable non-transcribed regions, and normally have faster rates of evolution

(Zarlenga & Higgins 2001; Perkins et al. 2011). However, these are the same gene cluster

as 18S rRNA, and so the difficulties of using a single marker remain. A demonstrative

figure is presented (Figure 1.7).

1.5 Objectives

Improving the current knowledge on distribution, diversity and evolution of

parasites is necessary. Apicomplexan parasites have been successfully amplified from

various hosts in CIBIO, including Hepatozoon species, using 18S rRNA specific primers

(Harris et al. 2012; Maia et al. 2012b; Tomé et al. 2012; Harris et al. 2013a). This allows

the understanding and estimating phylogenetic relationships of these parasites.

Apicomplexans diversity is huge and apart of certain groups with medical, veterinary and

economic interest is still poorly studied. The phylogenetic relations of the group are also

poorly studied. Reptiles, mammals and birds are some of the vertebrates hosts studied for

these parasites. In this study we focus on amphibians as hosts, perhaps the less explored

vertebrate group.

Amphibians can be used by these parasites either as unique intermediate host, or

as a first intermediate host later ingested by a second one. Therefore a wide diversity of

parasites is expected. In the case of Hepatozoon are the lineages found in amphibians

used as a first intermediate host closely related to those found in the predator? Or if the

parasite only uses amphibians as vertebrate hosts we can expect to observe a strong

coevolution. Many questions can be raised, for example, can we find the same parasite

lineages infecting different species or are they host-specific? Can we find mixed

Figure 1.7 – Diagram representing the organization of nuclear genes of ribosomal subunits. Scheme by José Babo.

30

infections, and different lineages in the same amphibian species? What is the intensity of

the different lineages, and what are the most common ones?

Considering all the different questions that can be made the objectives of this

thesis were:

1. Screen tissue and blood samples from different amphibian species

using specific parasite primers;

2. Assess the diversity of these parasites, specifically Hepatozoon;

3. Conduct microscopic surveys on available blood smears from these

hosts and compare detection with molecular methods;

4. Reconstruct the phylogenetic relationships of the sequences produced.

1.6 Organization of the thesis

Chapter 1 – General Introduction, Objectives

Introductory chapter with past studies and relevant information to contextualize the work

developed, as well as the general objectives and aims of this work.

Chapter 2 – Materials and Methods

Description of the techniques used during this work. Details of every step will be given,

procedures for sample collection to sequence analysis, passing trough DNA extraction

and blood smears.

Chapter 3 – Screening for Apicomplexan parasites in amphibians

This chapter presents the finding of a Dactylosoma ranarum in the amphibian Pelophylax

perezi using molecular methods, and the finding of several hemoparasites through

microscopic examination of blood smears.

Chapter 4 – Putative Ichthyophthirius identified in the amphibian Bufo calamita

through molecular screening.

This chapter presents the finding of an Ichthyophthirius in the amphibian Bufo calamita.

Chapter 5 - General Discussion

A general discussion of the results.

31

Chapter 6 – Final Remarks

This chapter presents the main conclusion of the study and ideas for future work. Apendix 1 Apendix 2

1.7 Taxonomy

Amphibian taxonomy around the regions targeted in this thesis has been suffering

some changes, and are in a state of flux. Therefore is this thesis we follow Beukema et al.

(2013), with the exception of one species, Epidalea calamita, which is going to be call

Bufo calamita to correspond to the same taxonomy used for the publication of one of the

works this thesis produced.

32

2 Chapter two

MATERIALS AND METHODS

33

2.1 Sample collection

A net was used to make the captures. After capturing the animals, a toe clip was

taken and the animal was released in the same place it was captured. Tadpoles were also

collected. In this case the entire animal was collected. The tissue samples were preserved

in 96% ethanol. If the cut in the toe resulted in bleeding, the blood was used to prepare

blood smears for microscopic observation and also stored in Whatman filter paper; for

detailed information see Appendix 1. GPS coordinates of the sampling location and the

date of capture were registered. Photographs were taken of each individual. Some

supplementary information (such as sex, age and size) was recorded when possible and

the presence of ectoparasites (e.g., ticks and mites) was checked and recorded. When

present, these were also collected. Host and parasite DNA extraction is possible from

both blood in Whatman filter paper and tissue samples.

The total number of samples used in this thesis was 353, from which 230 were

tissue samples, 72 blood samples and 51 slides. These samples are from a total of 269

individuals. The samples were collected prior to the beginning of this thesis. They include

12 species (Figure 2.2, Table 2.1), from locations spread throughout a vast area, including

the Iberian Peninsula, Mediterranean Islands, Macaronesia region and Morocco (Figure

2.1).

Figure 2.1 – Geographic distribution of the species used as samples.

34

Table 2.1 – List of amphibian host species used in this thesis.

Species Number of

samples

Bufo bufo 5

Amietophrynus mauritanicus 38

Discoglossus sardus 1

Bufo calamita 66

Hyla meridionalis 11

Hyla sarda 2

Pelobates cultripes 45

Pelophylax perezi 24

Pelophylax saharicus 82

Pleurodeles waltl 12

Bufotes balearicus 5

Bufotes boulengeri 11

Total 302

2.2 DNA extraction, PCR and Electrophoresis

The extracted DNA used for this thesis comes from two biological sources, blood and

tissue. The tissue was preserved in 96% ethanol and the blood was preserved in

Whatman filter paper and stored in the freezer at -20ºC. The DNA was extracted

employing the high salt extraction method (Sambrook et al. 1989). First a small portion of

tissue is cut is very small pieces, or a small portion of paper in the case of blood samples;

and collected into an eppendorf tube. Proteinase K (8 µl at 25 mg/ml) and lysis buffer (600

µl of solution composed of: 0,5M tris; 0,1M EDTA; 2% SDS; pH 8,0) are added to the

eppendorf; this allows the digestion of the tissue and releases the cellular contents. Heat

is also applied to help the reaction occur (typically 56º C). Ammonium acetate (300 µl of

solution composed of: 7M; pH 8,0) is used to precipitate the proteins, the sample is then

subjected to centrifugation (for 15 minutes at 14000 rpm at 0º C), and the supernatant

collected to a new eppendorf tube. The next step is to add ice cold isopropanol (600 µl),

used to precipitate the DNA into a pellet during centrifugation (for 25 minutes at 14000

rpm at 0º C). After, the DNA pellet is washed with ice cold 70% ethanol (1000 µl and

centrifuged for 15 minutes at 14000 rpm at 0º C) and left at room temperature, to allow the

35

Figure 2.2 –Photos of the species used during this thesis. A –Bufo bufo; B –Amietophrynus mauritanicus; C –Discoglossus

sardus; D –Bufo calamita; E –Hyla meridionalis; F –Hyla sarda; G –Pelobates cultripes; H –Pelophylax perezi; I –Pelophylax

saharicus; J –Pleurodeles waltl; K –Bufotes balearicus; L –Bufotes boulengerie. Photos E and I were taken by Daniele Salvi,

Photos A, C, D, F, G, H, J, K, L were taken by Matt Wilson, Photo B was taken by Pierre-Yves Vaucher.

ethanol to evaporate. The last step is to add ultrapure water (50 µl) to hydrate the DNA.

The full details for the method can be found in Appendix 2.

Several primer sets that were previously used for detection of multiple parasites

from host tissue samples, with satisfactory results at CIBIO producing sequences

matching apicomplexan parasites,(Harris et al. 2011; Maia et al. 2011; Harris et al. 2012;

Maia et al. 2012b; Tomé et al. 2012) were tested in amphibian hosts. The primers used in

this thesis for parasite screening were the pairs HepF300/HepR900 (Ujvari et al. 2004)

and HEMO1/HEMO2 (Perkins & Keller 2001), designed to target the 18S rRNA gene

region (Table 2). The Hep primers result in amplification product of about 600bp long,

while the HEMO primers amplify a fragment with about 900 bp long. Another pair of

primers were also used during this thesis, the CR-1 and CR-2 (Ellis et al. 1998). These

primers were design to amplify a fragment with around 900 bp of the 28S rRNA gene of

Coccidia (Table 2.2).

E F G H

I J K L

A B C D

36

Conditions and protocols relative to PCRs are described in Table 2.3, Table 2.4,

Table 2.5 and Table 2.6 for each pair of primers. PCRs were performed on a Biometra

TProfessional Standard gradient Thermocycler. Negative and positive controls were run

with each reaction. A positive control is a sample previously known to be infected and

negative control is the mixture of all reagents except the DNA. This is used to ensure that

the PCR reaction was successful and that no contaminations occurred. The PCR reagents

were all prepared and distributed in equal proportions by the reaction wells prior to the

adding of DNA. The DNA was added to each well separately. Two models of Taq were

used due to a changing of supply in the laboratory.

Table 2.2 – Details of the PCR primers used in this thesis.

Name Gene Sequence (5'→3') Reference

Hep300F 18S rRNA

GTTTCTGACCTATCAGCTTTCGACG Ujvari et al., 2004

Hep900R CAAATCTAAGAATTTCACCTCTGAC

HEMO1 18S rRNA

TATTGGTTTTAAGAACTAATTTTATGATTG Perkins & Keller 2001 HEMO2 CTTCTCCTTCCTTTAAGTGATAAGGTTCAC

CR1 28S rRNA

CTGAAATTGCTGAAAAGGAA Ellis et al., 1998

CR2 CCAGCTACTAGATGGTTCGA

Table 2.3 – Reagents used, and respective concentrations for the Hep, HEMO and CR primers, using invitrogen Taq DNA Polymerase.

The PCR products were run in in 2% agarose, containing 1 μl of GelRed Nucleic

Acid Stain (10,000x in water, BIOTIUM) per 50 μl of agarose solution. The gel is placed

Reagent Concentration

Hep HEMO CR

Water To 20 μl To 20 μl To 20 μl

Buffer 1 X 1 X 1 X

MgCl2 1.5 mM 3.75 mM 3.75 mM

dNTPs 0.125 mM each 0.2 mM each 0.2 mM each

BSA 0.4 mg/μl 0.4 mg/μl 0.4 mg/μl

Primer forward 0.6 μM 0.5 μM 1 μM

Primer reverse 0.6 μM 0.5 μM 1 μM

Taq 1 U 0.5 U 0.5 U

DNA 2 μl 2 μl 2 μl

37

into a mold and, after it solidified, was loaded. To load the gel a mixture of 2 μl of PCR

product and 2 μl of methylene blue solution was used for each well (the mixture was no

longer necessary when the PCR reaction was conducted using Bioline MyTaqTM DNA

Polymerase). A reference ladder was also loaded, to allow verification of the size of the

amplicons. Then the gel was placed in the electrophoresis equipment and run at 250

volts. Once the electrophoresis was completed, the results were checked using an

ultraviolet transilluminator and a photograph of each gel was taken.

Table 2.4 – PCR protocols for the Hep, HEMO and CR primers using invitrogen Taq DNA Polymerase.

Hep Hemo

CR

Step Function ºC Time Cycles ºC Time Cycles ºC Time Cycles

Initial Denature Denature 94 3’ 1 94 3’ 1 94 7’ 1

Thermal cycling

Denature 94 30’’

35

94 30’’

35

94 1’

35 Anneal 60 30’’ 48 30’’ 55 30’’

Extend 72 1’ 72 1’ 72 1’

Final Extend Extend 72 10’ 1 72 10’ 1 72 10’ 1

Hold Hold 10 ∞ 1 10 ∞ 1 10 ∞ 1

2.3 Sequencing and Data Analysis

The positive PCR products were sent to a private company (Macrogen Inc.), to be

purified and sequenced. Then the resulting chromatograms were checked manually and

assembled in BioEdit (Hall 1995). After that the sequences were blasted to identify closest

matches in GenBank. Two sequences matched with parasites.

Two methods were used to perform phylogenetic analysis. Maximum Likelihood

(ML), and Bayesian Inference (BI). Maximum Likelihood (ML) was performed using

PhyML 3.0 (Guindon et al. 2010), and Bayesian analysis was implemented using Mr.

Bayes 3.1 (Huelsenbeck & Ronquist 2001). The best fitting models of nucleotide

substitution for each alignment were chosen according to the Akaike Information Criteria

carried out in jModeltest 0.1.1 (Posada 2008).

38

Table 2.5 – Reagents used, and respective concentrations for the Hep, HEMO and CR primers, using Bioline MyTaq

TM DNA Polymerase.

Reagent Concentration

Hep HEMO CR

Water To 20 μl To 20 μl To 20 μl

Buffer 5 X 5 X 5 X

Primer forward 0.6 μM 0.5 μM 1 μM

Primer reverse 0.6 μM 0.5 μM 1 μM

Taq 1 U 0.5 U 0.5 U

DNA 2 μl 2 μl 2 μl

Table 2.6 – PCR protocols for the Hep, HEMO and CR primers using Bioline MyTaq

TM DNA Polymerase.

Hep Hemo

CR

Step Function ºC Time Cycles ºC Time Cycles ºC Time Cycles

Initial Denature Denature 95 3’ 1 95 3’ 1 95 7’ 1

Thermal cycling

Denature 95 30’’

35

95 30’’

35

95 1’

35 Anneal 60 30’’ 48 30’’ 55 30’’

Extend 72 1’ 72 1’ 72 1’

Final Extend Extend 72 10’ 1 72 10’ 1 72 10’ 1

Hold Hold 10 ∞ 1 10 ∞ 1 10 ∞ 1

2.4 Blood Smears

Whenever natural bleeding occurs blood smears were prepared. A drop of blood is

placed in a slide and with help from another slide (called the “spreader”) is spread. The

“spreader” should be placed at a 45° angle and backing into the drop. When the drop is

reached the blood should spread by capillarity across the edge which is then pushed

across the other slide. The slide is then left to be air-dried. Information about the sample

should be written at the end of the slide, preferably with a pencil. The “spreader” will

become the next slide to receive a smear, thus serving two purposes. Each slide can

accommodate more than one smear with proper identification.

39

The next step is fixing the smear, for that 100% methanol is used. Recipients that

can store slides in individual slots so they do not touch each other should be used. The

slides are immersed in 100% methanol for around 2 minutes. After being fixed, the slide

needs to be stained. The solution used for staining contains Giemsa and distilled water

with a pH around 7.0. The slides are immersed for a period of 50-55 minutes, followed by

a 2 minute wash with distilled water. Finally the smear is left to dry at room temperature

and is ready to be stored.

40

3 Chapter three

Screening for Apicomplexan parasites in amphibians

Article submitted 2013 to Herpetozoa

41

Screening for Apicomplexan parasites in amphibians

J. Seabra-Babo1, J.P. Maia2,3 D. James Harris2,3

1Departamento de Biologia, Universidade de Aveiro, Campus Universitário de Santiago,

3810-193 Aveiro, Portugal.

2CIBIO-UP, Centro de Investigação em Biodiversidade e Recursos Genéticos da

Universidade do Porto, Campus Agrário de Vairão, 4485-661 Vairão, Portugal

3Departamento de Biologia, Faculdade de Ciências, Universidade do Porto, Rua do

Campo Alegre FC4, 4169-007 Porto, Portugal

Author for correspondence:

james@cibio.up.pt

Introduction

Parasites are ubiquitous, but also a poorly known component of biodiversity, with

estimates of 0.1% of species described for some groups (Morrison 2009), while other

scientists simply note “we have no credible way of estimating how many parasitic

protozoa … exist” (Dobson et al. 2008). Yet parasites have dual interests for conservation

biologists, both for their impact on hosts with parasite-driven declines in wildlife becoming

increasingly common (Pedersen & Fenton 2007), but also because this relationship with

the hosts increases their risk of co-extinction, with the loss of hosts causing a cascading

extinction effect. Indeed, models suggest co-extinction may be the most common form of

biodiversity loss (Dunn et al. 2009). Given the difficulties in alpha-taxonomy of most

groups, and the lack of parasitologists (Šlapeta 2013), molecular screening, much like the

common “DNA barcoding” approach, may be an extremely valuable first assessment for

some parasites. Clearly, integrative approaches combining morphological and molecular

data would be preferable given time (Will et al. 2005), but molecular screening has many

advantages, besides being relatively quick and easy. For a start, sequence data is

invaluable for placing parasites in a phylogenetic framework. Furthermore, many times

host samples are collected for other purposes, such as host genetic assessments, but

these same samples can be reused in parasite screening studies.

42

Apicomplexan blood parasites are a prime example of a group where screening

can be useful. Assessments of these have in the past focused on groups with strong

anthropogenic interests due to health reasons, such as Plasmodium, or groups with

significant economic impact. Others, such as Hepatozoon, despite being the most

common blood parasite of reptiles (Telford 2009) gained little attention. Molecular

screening however, using primers specific for a section of the 18S rRNA gene, has greatly

clarified phylogeny (e.g. Barta et al. 2012; Harris et al. 2012) identified infections in new

host orders (e.g. Pinto et al. 2012) and indicated that predator-prey trophic pathways may

be widespread in some cases, such as between lizards and snakes (Tomé et al. 2013). At

the same time, detection of other parasites such as Stramenopiles (Maia et al. 2012a),

gives further valuable information. Despite this, screening with specific primers can be

misleading if some parasites are not detected (Zehtindjiev et al. 2012). It is therefore

necessary to test this aspect in different host groups.

Amphibians as a whole have suffered global declines, and parasites are a key

driving factor (Beebee & Griffiths 2005). Although the role of the fungus Batrachochytrium

dendrobatidis is widely accepted (e.g. Daszak et al. 2003), screening for other parasites is

needed. For example, Hepatozoon were first recorded in one of the three amphibian

orders, Gymnophiona, only recently (Harris et al. in press). Various Hepatozoon have

been identified in amphibians using microscopy (e.g. Stenberg & Bowerman 2010), and

they were detected during screening of an introduced population of frogs, Pelophylax

perezi, from the Azores islands (Harris et al. 2013c). On the other hand, a screening of

Bufo calamita from the Iberian Peninsula did not detect Apicomplexan parasites, only a

putative Ichthyopthirius multifiliis, an important ciliate pathogen of fish (Harris et al.

2013b).

The aim of this study was to screen a large number of amphibians from Europe

and North Africa, using samples that had been primarily collected for studies of the host.

Because of this, in most cases blood smears to assess prevalence under the microscope

were not available. However, the same technique had proved effective in amphibians from

the Azores, and in reptiles from this region (e.g. Maia et al. 2012b). Thus we had prior

reasons to expect the method to be effective. Since it is known that these primers can

detect other organisms, all positive samples were sequenced. When few positives were

identified (see results), additional samples were collected specifically for both molecular

screening and microscopy. These were then compared to assess the efficiency of

detection using molecular screening.

43

Materials and Methods

Tissue samples (toe clips) were collected from 136 amphibians belonging to 6

species, from various localities in the Iberian Peninsula, the Balearic islands and Morocco

and stored in 96% ethanol (Table 1). The taxonomy of many amphibians in this region is

in a state of flux, but here we follow Beukema et al. (2013). For a smaller number of

specimens, blood drops stored on Whatman paper (56 specimens, 4 species) and blood

smears (51 specimens, 4 species) were also available (indicated in Table 4.1). Blood

smears were stained with Giemsa as described elsewhere (e.g., Telford, 2009) and

examined using an Olympus CX41 microscope with an in-built digital camera (SC30)

(Olympus, Hamburg, Germany). Several photomicrographs per slide were taken at 400×

magnification and stitched using cell^B software (basic image-acquisition and archiving

software, Olympus, Münster, Germany). When no parasites were identified after circa 10

minutes of examination, the slides were considered negative. When parasites were

identified, even in very low numbers, slides were scored as positive. For some examples,

intensity of infection was estimated based on numbers of parasites per 3,000 cells.

Table 4.1 –Amphibian host species screened for parasites, the number tested using alternative

source material (tissue or blood), and the number examined under the microscope on slides.

DNA was extracted using standard High Salt methods (Sambrook et al. 1989).

Detection of blood parasites was made using PCR reactions with the primers HepF300

and HepR900 (Ujvari et al. 2004), which were designed to amplify Hepatozoon parasites.

Conditions of the PCR are detailed in Harris et al. (2011). A limited subset (47) of samples

was also tested with the Hemo1 and Hemo2 primers (Perkins & Keller 2001). Although

Species Tissue Blood Slides

Pelobates cultripes 45 - -

Pelophylax saharicus 52 30 26

Hyla meridionalis - 11 7

Amietophrynus

mauritanicus 30 8 10

Bufotes boulengeri 4 7 8

Bufo bufo 5 - -

44

these are known to be less efficient at detecting Hepatozoon relative to the Hep primers

(Maia et al. 2012b), they were nonetheless tested in case they gave results in amphibians.

Negative and positive controls were run with each reaction. PCR products were analyzed

by electrophoresis in 2% agarose and visualized by Gel Red staining and UV

transilumination. The positive PCR products obtained were purified and sequenced by a

commercial sequencing facility (Macrogen Europe, The Netherlands). Positive samples

can be caused by contaminants, such as fungi (Tomé et al. 2012). Thus all positive PCRs

were compared against the public database GenBank, using a BLAST similarity search.

Results

Initial screening was carried out on the toe clips, as these are most readily

available from genetic studies of the vertebrate hosts. Only a single positive sample for

Apicomplexan parasites was found with the Hep primers, and the BLAST comparison

showed a 99.8% similarity with Dactylosoma ranarum from a Pelophylax kl esculentus

host from Corsica, France (Accession numbers HQ224957 and HQ224958, (Barta et al.

2012). Only a single nucleotide differed from the GenBank sequences over 613bp of

compared sequence data. Analysis of blood samples also failed to detect any positive

infections, as did PCRs conducted with the Hemo primers. However, when blood slides

were examined various positive samples were identified (Figure 4.1). These were found in

two different host species (Pelophylax saharicus and Amietophrynus mauritanicus), and at

least for P. saharicus infection rates were high (12 in 26 individuals screened). Based on

intensity infections, the positive for Dactylosoma had the lowest parasitaemia level (0.1%

infected cells), but was still detected using the molecular method, while intensities of

apparent Hepatozoon infection, ranged up to 3% in the sample of A. mauritanicus

(DB15569), but which scored as a negative in the molecular screening.

Discussion

Screening for parasites using conserved primers has the potential to greatly

improve knowledge on parasite diversity and distribution. Like all molecular approaches, it

can be improved by adoption of an “integrated” approach, as barcoding approaches can

be misleading in certain circumstances (Will et al. 2005). However, given that hundreds of

45

samples can be quickly assessed to give a rapid first overview of parasite prevalence, and

at the same time can sometimes detect unexpected forms, screening is likely to become

more and more common in the future. Given this, it becomes essential to know which

primers can be used in different circumstances.

For most studies of Hepatozoon, the Hep primers (Ujvari et al. 2004) have proven

to be efficient at detecting not only divergent Hepatozoon lineages (e.g. Harris et al.

2012), but also various other parasites. In amphibians they have detected at least two

unrelated lineages, from the frog host Pelophylax perezi and from caecilians of the genus

Grandisonia (Harris et al. 2013c, in press respectively). Most studies have indicated that

identification efficiency was at least as high, or even higher, when compared to

assessment of blood smears (e.g. O’Dwyer et al. 2013). Yet in this study they failed to

amplify any Hepatozoon, which were clearly identified in at least two of the species

screened. One possibility is that toe clips are not ideal sources of material for studies of

Figure 2.1 –Images of positive Hepatozoon infections in A) A. mauritanicus, and B) P.

saharicus, both of which failed to amplify using the screening protocol employed, C) a positive

infection of presumed Dactylosoma ranarum in Pelophylax saharicus, and D) a typical negative

sample. The scale bar corresponds to 20μm.

46

these parasites in amphibians. However, we also did not detect any parasites in blood

drops stored in Whatman paper, which implies that the source of the host sample was not

the issue. Rather the primers used failed to amplify Hepatozoon lineages in these hosts.

This is unexpected, given that they have amplified Hepatozoon from P. perezi in the

Azores islands, but not P. perezi or the related P. saharicus in the Iberian Peninsula and

N. Africa. On the other hand, it is clear that many different Hepatozoon lineages can be

found in the same intermediate host species (e.g. Tomé et al. 2013). It is possible that

some common Hepatozoon are not detected with these primers, while other lineages

which occasionally occur are amplified. Parasites can sometimes be overlooked, in both

molecular screening and under the microscope, when parasitaemia levels are very low.

This might explain why we detected Dactylosoma but not Hepatozoon, if parasitaemia

levels were much lower in the latter. However, our assessment of parasitaemia levels

indicates the opposite, with a very low level for Dactylosoma (just 0.1% of erythrocytes

infected, an order of magnitude lower than some infection rates with Hepatozoon). The

identity of the Hepatozoon identified under the microscope remains unclear. Forty two

Hepatozoon have been associated with amphibian hosts (Smith 1996), and since in much

of the earlier literature the hosts were not identified below the generic level, previous

identifications of Hepatozoon from toads and frogs from North Africa may well correspond

to P. saharicus and A. mauritanicus.

The finding of Dactylosoma in one sample of P. saharicus increases the list of

parasites that have now been detected using molecular screening with these primers. The

very high similarity with the sample from GenBank indicates it is very likely to be D.

ranarum, and implies low intermediate host specificity for this parasite. Indeed, D.

ranarum is thought to occur in amphibians from Africa and North and South America, Asia

and Europe (Barta 1991). Genetic data from hosts from other regions will be invaluable in

assessing this further.

To conclude, although these primers have been used in screening various

mammal and reptile hosts for Hepatozoon, they do not appear to be useful for

amphibians. However, they did detect Dactylosoma, and may be useful for screening for

these parasites. Hepatozoon were detected in P. saharicus and A. mauritanicus in

Morocco using microscopy. New primers need to be developed to amplify in these

amphibian parasites, so that they can be appropriately identified and placed in a

phylogenetic framework.

47

Acknowledgements

This work forms part of the MSc thesis of J.S-B., supervised by DJH. JPMCM is

supported by a Fundação para a Ciência e a Tecnologia (FCT) PhD grant

(SFRH/BD/74305/2010) and co-financed by FSE and POPH and EU. DJH is partially

supported by FEDER through the COMPETE program and the Project “Genomics and

Evolutionary Biology” cofinanced by North Portugal Regional Operational Programme

2007/2013 (ON.2 – O Novo Norte), under the National Strategic Reference Framework

(NSRF), through the European Regional Development Fund (ERDF). Fieldwork in North

Africa was supported by the Percy Sladen fund (to DJH) and by the Chicago

Herpetological Society (to JPMCM). The authors are grateful to their many colleagues

who contributed samples for this study.

References

The bibliography used in this chapter can be found in the References chapter of

this thesis

48

4 Chapter four

Putative Ichthyophthirius identified in the amphibian Bufo calamita through

molecular screening.

Article published in 2013 in BULLETIN OF THE EUROPEAN ASSOCIATION

OF FISH PATHOLOGISTS 33 (1), 24-27)

49

Note

Putative Ichthyophthirius identified in the amphibian Bufo calamita through

molecular screening.

D. J. Harris1 2, J. Seabra-Babo4, J. Tavares4 and J. P. M. C. Maia1 2 3

1CIBIO-UP, Centro de Investigação em Biodiversidade e Recursos Genéticos da

Universidade do Porto, Campus Agrário de Vairão, 4485-661 Vairão, Portugal;

2Departamento de Biologia, Faculdade de Ciências, Universidade do Porto, Rua

do Campo Alegre FC4, 4169-007 Porto, Portugal;

3Institut de Biologia Evolutiva (CSIC-UPF). Passeig Marítim de la Barceloneta, 37-

49. 08003 Barcelona. Spain;

4Departamento de Biologia, Universidade de Aveiro, Campus Universitário de

Santiago, 3810-193 Aveiro, Portugal.

Abstract

The protozoan parasite Ichthyopthirius multifiliis is an important pathogen of many

fish species. During molecular screening of 18S rRNA gene, we identified an apparent

Ichthyopthirius from the amphibian Bufo calamita, to our knowledge the first such

detection reported from a wild amphibian. This has important implications for parasite

control approaches in aquaculture.

Protistans are one of the most important groups of pathogens of fish (Scholz

1999). Of these, one of the most pathogenic protozoan parasites of freshwater fish is the

ciliate Ichthyophthirius multifiliis, which causes Ichthyopthiriasis, or “white spot disease”

(Dickerson 2006). This is a major problem in aquaculture, compounded by its

cosmopolitan distribution, having been reported from all regions where fish are cultivated.

Furthermore, it affects a wide range of fish including carp, trout, eel, catfish and

ornamental fish (Scholz 1999). Water temperature appears to be critical for disease

outbreaks, which are more common when fish are stressed and water temperature rises

(Dickerson 2006). One important, unanswered question is how does an obligate parasite

survive between outbreaks? It has been hypothesized that it is most likely that the

parasite is maintained through low level infections (Dickerson 2006).

50

Although identification of parasites was traditionally based on morphological

characters, molecular techniques are a powerful tool to identify various parasite groups in

freshwater environments. Not only can larger numbers of samples be screened quickly,

but molecular data can also give information regarding the diversity of parasites and their

phylogenetic relationships.

During routine screening of the amphibian Bufo calamita from the Iberian

Peninsula a standard protocol was followed. DNA was extracted from toe-clips using

standard high-salt methods (Sambrook et al. 1989). Primers used were HepF300 and

HepR900 (Ujvari et al. 2004), that target a part of the 18S rRNA gene. They are known to

amplify Apicomplexan parasites (Harris et al. 2012) as well as various other groups

including Stramenopiles (Maia et al. 2012a) and some fungi (Tomé et al. 2012) .PCR

conditions consisted of 35 cycles of 94ºC (30 seconds), 60ºC (30 seconds) and 72ºC (1

minute) – see Harris et al. (2011)for more details. Negative and positive controls were run

with each reaction. All positive PCRs were sequenced by a commercial service

(Macrogen Inc.). Resulting chromatograms were checked manually and assembled in

BioEdit (Hall 1995). BLAST was used to identify closest matches in GenBank.

Out of the 56 samples of B. calamita screened (10 from Castelo Branco, 5 from

Mindelo and 19 from Aveiro in Portugal, and 22 from Trabazos in Spain), two positives

(DB16921 and DB16925 from Trabazos) matched ciliates in the BLAST search (GenBank

Accession numbers KC512767 and KC512768 respectively). The first gave the closest

match with Hausmanniella discoidea (Accession number EU039900, Dunthorne et al.,

2008). The match was 98% with 7 identified differences over 531bp. Hausmanniella are

widespread protozoans, often identified in environmental assessments (e.g. Bartosŏva

and Tirjakova (2008)). However, the second ciliate gave a closest match with I. multifiliis

(Accession number U17354, Wright and Lynn (1995)). To confirm this assessment, two

different phylogenetic analyses were carried out (Maximum Likelihood and Bayesian

Inference) using the most closely related sequences available on GenBank. Maximum

Likelihood (ML) analysis included random sequence addition (100 replicate heuristic

searches), and support for nodes was estimated using the bootstrap technique

(Felsenstein 1985) with 500 replicates, using PhyML 3.0 (Guindon et al. 2010). The AIC

criteria carried out in jModeltest 0.1.1 (Posada 2008) were used to choose the model of

evolution employed (TrN+G). Bayesian analysis was implemented using Mr. Bayes 3.1

(Huelsenbeck & Ronquist 2001) with parameters estimated as part of the analysis. The

analysis was run for 5x106 generations, saving one tree each 1000 generations. The log-

likelihood values of the sample point were plotted against the generation time and all the

51

trees prior to reaching stationary were discarded. Remaining trees were combined in a

50% majority consensus tree. Following , Paramecium tetraurelia (X03772) was

designated as an outgroup (Figure 3.1).

As expected, the ciliate identified from B. calamita was sister taxa to I. multifiliis with

high support. Sister taxa to this was Ophryoglena catenula (U17355, Wright and Lynn

(1995)), then two uncultured clones (EF586162 and EF586110,Dopheide et al. (2008)),

and then Tetrahymena paravorax (EF070253,Chantangsi et al. (2007)), relationships

similar to those previously proposed (Wright & Lynn 1995).

This result is important for several reasons. First, I. multifiliis is currently

considered to exclusively parasitise freshwater fish, although tadpole stages of the marsh

frog Limnodynastes peronii were successfully infected experimentally (Gleeson 1999).

Our results indicate that the parasite DNA could be detected from the clipped toe of a wild

amphibian. This does not per se demonstrate infection - during the tomont stage of the life

cycle of I. multifiliis it produces a sticky capsule that can attach to various substrates.

From this capsule theronts are released which can survive for 2-4 days while actively

searching for a host. Whether detection indicates an infection or not, clearly needs further

investigation. Secondly the sequence shows limited differentiation from the published I.

Figure 3.1 – ML tree of the apparent Ichthyophthirius from a Bufo calamita, and closest available comparative sequences from GenBank. Support for the Bayesian and for ML analysis are given above and below the nodes, respectively. The branch of Paramecium tetrawelia was shortened 75%.

52

multifiliis sequence, with 12 differences over 544 bp. It has previously been proposed that

multiple strains or even cryptic species may occur within I. multifiliis (Nigrelli et al.

1976).and our results seem to be further evidence for this with the two sequences

showing some divergence whilst clearly belonging to the same clade.

Given the importance of I. multifiliis as a parasite of freshwater fish, it becomes

imperative to ascertain if other amphibian species can also host similar parasites, and

whether these can cross infect between amphibians and fish. If amphibians are acting as

sinks for these parasites, it will be important to take this into account when trying to control

disease outbreaks. Further molecular screening of additional fish and amphibian species,

either using general primers as in this study or ones more specific for Ichthyopthirius, will

be essential to provide further information on this possibility.

JPMCM is supported by a Fundação para a Ciência e a Tecnologia (FCT) PhD

grant (SFRH/ BD/74305/2010) and co-financed by FSE and POPH and EU. Thanks to the

anonymous reviewer for their useful comments on an earlier version.

References

The bibliography used in this chapter can be found in the References chapter of

this thesis.

53

5 Chapter five

General discussion

54

General Discussion

In this chapter I will present a general discussion of the work performed. As the

two previous chapters present the papers produced during this thesis, this chapter will be

dedicated to an overall evaluation of the results. Possible explanations for the results will

be suggested, and an attempt to clarify the reasons for choosing molecular methods as

the main tool.

In the beginning of the work for this thesis we proposed to screen several tissue

samples of amphibians, with the objective of detecting parasites with a focus on

Hepatozoon species. For that 230 samples from 8 species were subjected to PCR using

Hep primers (Ujvari et al. 2004). The samples came from several locations, this would

have permitted to analyse differences between lineages of parasites detected, and also

what lineages were most common and differences between intensities. A smaller subset

of samples (47) were also tested with the Hemo1 and Hemo2 primers (Perkins & Keller

2001) and CR1 and CR2 primers (Ellis et al. 1998).

The primary use of Hep primers (Ujvari et al. 2004) come from the previous

success they demonstrated, detecting Hepatozoon in several other studies (Harris et al.

2012; Maia et al. 2012b; Tomé et al. 2012; Harris et al. 2013c). Hemo primers, although

less efficient than Hep for other groups of vertebrates (Maia et al. 2012b), were tested in

case they gave results in amphibians. The use of CR primers was justified by the

possibility of detecting other groups of parasites. The entire sample set tested with Hemo

and CR primers was negative. Nonetheless PCR reactions with Hep primers produced 27

positives. All these positives were sequenced, and since contaminants or other false

positives can be the cause of them, sequences were compared against the public

database GenBank, using a BLAST. Blast search results in only two parasites with the

majority of sequences corresponding to fungi. Fungi have been reported in other studies

with these primers (Tomé et al. 2012). The two more interesting parasites were a putative

Ichthyopthirius multifiliis detected in Bufo calamita from Castelo Branco, Portugal, and a

Dactylosoma ranarum in a Pelophylax saharicus from Morocco.

Although for fewer samples, other source of material for DNA extraction was

available, primarily blood drops from individuals that presented natural bleeding upon the

collection of the toe clip. Even though toe clips proved efficient in other studies (Harris et

al. 2013c), as different sources can influence the result of the PCR (McKenzie & Starks

2008), these samples were also tested. A total of 72 samples were available from 8

species. Again Hep primers were used, and a small number of samples (23) were tested

55

with Hemo and CR primers. All blood samples failed to detect any positive infections,

implying that the source of the material was not the problem.

With such results there were only two possible explanations: either all the

amphibian samples were free of Hepatozoon infections; or the primers fail to detect these

infections, perhaps because of genetic differences to Hepatozoon parasites found in other

vertebrate groups. Several blood smears stained with Giemsa were collected, and the

answers to the possibilities mention above reside in the observation of these smears. Fifty

one slides were available from 4 species and all were examined using using an Olympus

CX41 microscope with an in-built digital camera (SC30) (Olympus, Hamburg, Germany).

The slides were observed for 10 minutes and after, if no parasite was found they were

consider parasite free, however if parasites were observed, even in very low numbers the

slides were considered positive for infection. When positive the intensity of infection was

estimated based on numbers of parasites per 3,000 cells. From 51 slides 13 were

positive. The species with positive score were Amietophrynus mauritanicus, (10%, 1/ 10),

and Pelophylax saharicus, (46%, 12/26). In two slides hemogregarines were clearly

identified (DB15569 from Amietophrynus mauritanicus, DB14709 from Pelophylax

saharicus).

Despite the differences in sample size between the two species, an analysis to the

ecologic necessities of these species can give some clues about the differences in

prevalence. Regarding habitat, Amietophrynus mauritanicus can be encountered near

both permanent or temporary water bodies, while Pelophylax saharicus is more restricted

to permanent water bodies (Le Berre 1989; Chillasse et al. 2002). Differences are also

found regarding the feeding habits of these two species, A. mauritanicus feeds primarily of

insects, while P. saharicus feeds from insects, fishes and even other frogs (Doumergue

1901; Chillasse et al. 2002). The question is: Can these differences in feeding habits

explain the difference in prevalence numbers? Transmission by mosquito bite and trophic

transmission is known to occur in frogs (Ferguson & Smith 2012). Therefore, frogs can be

infected while serving as a meal for an infected mosquito, or while feeding from infected

preys. Taking this into account, the higher numbers presented by P. saharicus 40%

compared to A. mauritanicus, 10%, can be the reflection of infections transmitted by

insects plus the feeding of infected fishes and frogs. The apparent higher exposition to

potential parasites hosts can suggest that P. saharicus can be more susceptible to

parasite infections.

Although P. saharicus presented higher prevalence, that does not mean higher

level of pathogenicity. The impact of these parasites to their hosts can be significant,

56

however is still not determined in many species, and may vary according to the species.

Some parasite species can be tolerated with low level infections and cause severe illness

when high levels of infection is presented without being fatal for the host, while other are

highly pathogenic and result in death (Baneth 2011). Hepatozoon species that infect

amphibians form a separate clade, therefore can present unique characteristics and

implications for their hosts. Further investigation is required to assess how these parasites

are affecting amphibians and the implications of these associations.

The positive results using microscopy suggest that the primers used fail to detect

parasites. Although microscopy of gamonts is not adequate to identify the parasites to a

specific level, two identified as Hepatozoon sp. and several without classification, it

demonstrate a higher identification efficiency over PCR for these primers, at least for

amphibians samples.

This difficulty identifying hemogregarines based on morphological characters is the

main reason why the use of molecular tools has been increasing. The use of

morphological characters that are many times homoplastic in the past created a huge

controversy surrounding knowledge about parasite diversity and distribution. The lack of

information about some groups and the use of these characters to establish taxonomy

and phylogenetic relations have originated wrong taxonomic placements and phylogenies.

Although the use of molecular tools has been improving our knowledge about parasites, it

also sometimes creates conflicts with taxonomy accepted using traditional tools (Barta

2001). Molecular tools allow to process quickly large quantities of samples, and the ability

to distinguish individuals that possess similar morphological characters. However as seen

in this thesis to use molecular tools such as PCR some precautions have to be taken, and

selecting appropriated primers is fundamental.

Simply saying that the primers fail to detect Hepatozoon that were clearly visible

under the microscope is not the only explanation possible. Hep primers are known to

amplify several lineages of Hepatozoon even within the same intermediate host (Tomé et

al. 2013). Given that they were not able to detect infections in this thesis, it is possible that

the lineages identified using microscopy are different lineages from the previously

reported. The current information is not enough to establish if: either they are distant

lineages or uncommon ones, which make the primers used unable to detect them. These

hypotheses are interesting from an evolutionary perspective, and deserve attention for

future researchers. New primers and testing different genes is required to clarify these

possibilities.

57

6 Chapter six

Concluding remarks and future possibilities

58

6.1 Concluding remarks

The work developed for this thesis was an attempt to gather knowledge about

parasites from amphibians, particularly Hepatozoon. It was intended to be a large

molecular survey of the diversity and distribution of parasites in amphibians from the

Mediterranean region and North Africa. Not a single Hepatozoon infection was detected,

and only two key parasites were detected, what was unexpected since the primers used

presented good results with other samples (Harris et al. 2012; Maia et al. 2012b; Tomé et

al. 2012; Harris et al. 2013c). Nonetheless microscopy was able to detect several

infections with at least two apparently being Hepatozoon.

In the first study 56 individuals of the species Bufo calamita were screened for the

presence of parasites using specific primers, known to work with other vertebrates (Harris

et al. 2011; Maia et al. 2012b; Tomé et al. 2012). Two infections were detected in two

different individuals. Both infections appear to be ciliates, probably Hausmanniella

discoidea and Ichthyophthirius multifiliis. The first is often identified in environmental

assessments, however the second is considered to exclusively parasitize freshwater fish.

This was the first report of this parasite in a wild amphibian host. However, detection itself

may not reflect infection, since these parasites during the tomont stage of the life cycle

produce a sticky capsule that can attach to various substrates. The phylogenetic

relationships of this sequence revealed a similarity to those previously proposed for

Ichthyophthirius multifiliis (Wright & Lynn 1995), although limited differentiation from the

published sequence was showed. This suggests that multiples strains or even cryptic

species may occur within I. multifiliis, supporting the theory of Nigrelli et al. (1976).

Ichthyophthirius multifiliis is considered an obligate parasite and questions about

how the parasite survives between outbreaks are still unanswered. There has been

hypothesized that low level infections is used to maintain the parasite. However with a

finding like these further questions rise. Are amphibians acting as reservoirs for these

parasites between outbreaks? Can cross infections between amphibians and fish occur?

And what other species of amphibians are parasitized by this organism?

Before trying to answer these questions the infection should be confirmed. With

this, future work should aim at screening both fish and amphibians with these or more

specific primers. More variable markers than the one used (18S rRNA) would be

particularly valuable. The finding of this parasite in an amphibian sample should also be

taken into account when planning for disease outbreak control.

For the second study tissue and blood samples were used as well as blood

smears. 136 tissue samples from six different species of amphibians were used, 56 blood

59

drops stored in Whatman filter paper from 4 different species and 51 blood smears from 4

species also. The samples were tested with parasite specific primers and the blood slides

were examined using an Olympus CX41 microscope with an in-built digital camera.

Samples were scored as negative when no parasites were observed under the

microscope and positive when parasites were observed even in very low numbers. An

apicomplexan parasite however was detected, Dactylosoma ranarum from a Pelophylax

saharicus tissue sample. The parasite found show very high similarity with a comparable

sample from GenBank, which was reported in a Pelophylax kl esculentus, implying that

the parasite presents low intermediate host specificity. Concerning the blood smears

several slides were scored positive and two were identified as Hepatozoon. This implies

that the primers fail to detect these infections. This suggest that either these primers are

not efficient with amphibian samples or at least some of the Hepatozoon lineages of

amphibians are quite distant to the ones found in reptiles, such lizards and snakes, and so

did not amplify. Therefore new primers need to be developed to detect Hepatozoon in

amphibians. The finding of Dactylosoma using Hep primers increases the list of parasites

that have now been detected with them.

Hep primers fail to detect Hepatozoon infections in this thesis, nonetheless they

were able to detect an Apicomplexa parasite and proved efficient in other studies (Harris

et al. 2012; Maia et al. 2012b; Tomé et al. 2012; Harris et al. 2013c). As in other studies

(Harris et al. 2012; Maia et al. 2012a; Tomé et al. 2012), detection of other organisms also

occurred. This suggests that Hep primers are less specific than previously thought, which

could cause incorrect infection estimates. Therefore it is very important to validate the

PCR products amplification through the use of sequencing, comparing with published

sequences (e.g. BLAST), and performing phylogenetic analyses. Molecular screening

through PCR is a very useful tool with many advantages, nonetheless it also has

limitations as shown in this study. However, these limitations can be overcome by

optimization of the technique and development of more specific assays, and it should be

complemented with the use of traditional techniques, such as microscopy.

In general, the results obtained may lead to thinking that the used primers are not

effective for screening for hemoparasites in amphibians. However, the Hep primers allow

the detection of several different organisms, in this thesis two ciliates and an

apicomplexan. Unlike other studies (Richard et al. 2002; Durrant et al. 2006), microscopy

has proven more efficient than PCR in this thesis. This can be explained by the already

mentioned inefficiency of the primers. New primers need to be developed for routine

screen of Hepatozoon parasites in amphibians. Other methods of DNA extraction can also

60

be considered as a possible factor influencing the inefficacy of the PCR technique. Using

commercial kits or other sources of tissue can influence the outcome of the PCR.

Nonetheless these are just speculative assumptions, since the amplification of

Hepatozoon from infected amphibians was possible using commercial kit to extract DNA

from P. perezi from the Azores (Harris et al. 2013c). Trying to assess the diversity of

Hepatozoon in amphibians is required to increase the knowledge of this group. It also

would provide valuable information for phylogenetic and distribution studies. Much work is

yet to be done.

6.2 Future possibilities

The knowledge about parasite diversity is still poor. Improving the knowledge

about the Hepatozoon group will help to increase the information available to

parasitologists and will allow more accurate phylogenetic reconstructions. However more

investigation is necessary to improve this lack of information. Combining information about

different groups of hosts (snakes, lizards, amphibians, and mammals) will enhance the

overall view of the group. Also the use of samples from different location as well as from

different seasonal periods, will allow testing hypothesis about the development and

ecology of hemoparasites. Implementing new methods, like real-time PCR, which is not

only more accurate that the conventional PCR, but also allows to estimate the level of

parasitaemia, evaluate and compare differences among the type of tissue and DNA

extraction technique, factors that can influence the results.

Meanwhile, developing new primers and continue routinely screening of

amphibians parasites, both through microscopy and molecular tools, is required. More

sampling from different species and different regions are crucial. There is need to sample

a much broader range of taxa, not focusing the efforts solely on parasites of medical and

veterinary importance. Developing new protocols which use alternatives markers to the

18SrRNA, and generating multiple sequences from different genes is necessary to

improve the estimate of phylogenetic relationships and overall diversity.

Also very important is identifying the hematophagous invertebrate hosts to assess

complete life cycles for the species within the Hepatozoon group. Therefore collecting

ectoparasites is a necessity. In CIBIO a vast collection of these invertebrates has been

assemble over the years, ticks, mites, leeches and others. It is necessary to start

processing these samples, and continue to collect them during field work. Summing up,

future research should focus on improved sampling and optimization of the molecular

methods for routine parasite screen in amphibians

61

7 References

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Adl, S.M., Simpson, A.G., Farmer, M.A., Andersen, R.A., Anderson, O.R., Barta, J.R. et al. (2005). The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. Journal of Eukaryotic Microbiology, 52, 399-451.

Aikawa, M., Miller, L., Johnson, J. & Rabbege, J. (1978). Erythrocyte entry by malarial parasites. A moving junction between erythrocyte and parasite. The Journal of Cell Biology, 77, 72-82.

Aisien, M.S.O., Nago, S.G.A. & Rodel, M.O. (2011). Parasitic infections of amphibians in the Pendjari Biosphere Reserve, Benin. Afr. Zool., 46, 340-349.

Allen, K.E., Yabsley, M.J., Johnson, E.M., Reichard, M.V., Panciera, R.J., Ewing, S.A. et al. (2011). Novel Hepatozoon in Vertebrates From the Southern United States. The Journal of parasitology, 97, 648-653.

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Appendix 1

Extracting DNA from blood samples stored in Whatman filter paper is possible. For

this the samples should be stored like in the picture presented and the following steps

need to be done.

1. Identify the paper with sample information (location, date), preferably using

a pencil;

2. In each division place a some drops of blood;

3. Let the blood air dry;

4. Store the disk in a zip-lock plastic bag with some silica gel to prevent

damage by humidity. Push all the air out of the bag;

5. Preferably one disk should be store by bag, but if the space is limited

multiple disks can be stored in the same bag. For this blood drops need to

be already dry and a blank, clean filter paper disk is placed between papers

with blood drops;

6. The bags containing disks can be at ambient temperature until returning to

the lab, and then they should be stored in a freezer at -20ºC.

7. To extract the DNA remove the disks from the freezer and allow them to

reach room temperature.

Draw by José Babo

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Appendix 2

We can divide the High-salt method of extraction in 3 steps. First the preparation of

the material, second the addition of the reagents and finally preparation of DNA for

analysis. These three steps will be explained in detail.

1. First the bench should be cleaned with ethanol;

2. A glass also cleaned with ethanol should be marked in divisions in equal

number of samples to be extracted;

3. Mark the eppendorfs with the sample code;

4. Light up a lamp and prepare a cup with ethanol for the dissection material;

5. Use fire for sterilization of the material;

6. Cut the tissue in small pieces and put it in the corresponded eppendorf.

Sterilize the material between each sample;

After all these preparations, the next phase is the addition of the reagents.

1. First add 600 µl of Lysis buffer (0,5M tris; 0,1M EDTA; 2% SDS; pH 8,0) to

each eppendorf;

2. Using a pipette add 8 µl at 25 mg/ml of proteinase K, mix well with the

vortex and incubate overnight at 56ºC;

3. Put the tubes in the freezer for 30 minutes (for the solution to cool);

4. Add 300µl of ammonium acetate (7M; pH 8,0) to each tube, agitate and

centrifuge for 15 minutes at 14000 rpm at 0ºC. The precipitation of proteins

will form a white pellet at the bottom of the tube (if precipitated proteins

remain in the supernatant, add 100 µl of ammonium acetate and centrifuge

again);

5. Label new eppendorfs with the corresponding codes and transfer the

supernatant into these, add 600 µl of ice-cold isopropanol. This step will

allow the precipitation of DNA;

6. Leave the tubes in the freezer for 3hours to overnight;

7. Centrifuge for 10-30 minutes at 14000 rpm at 0ºC, put the eppendorfs in

the centrifuge with the opening lid turned to the centre, since the DNA

pellet will form in the other side. Discard the supernatant;

Finally the DNA is ready to be prepared to for analysis.

1. Pipette 1000 µl of ice-cold 70% ethanol to the tubes and mix well tapping

the end of the tube until the DNA pellet is released;

2. Centrifuge for 15 minutes at 14000 rpm at 0ºC and discard the supernatant;

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3. Leave the eppendorfs with the lid open at room temperature or in the

incubator for the ethanol to completely evaporate;

4. When the ethanol is completely evaporated add 50 to 200 µl of ultra-pure

water (or other DNA hydration solution) and leave to hydrate for at least 2

hours (preferably overnight) at room temperature with agitation.