Genetic subtypes in unicellular intestinal parasites with special focus on Blastocystis

Joakim Forsell

Department of Clinical Microbiology Umeå 2017 Responsible publisher under Swedish law: the Dean of the Medical Faculty This work is protected by the Swedish Copyright Legislation (Act 1960:729) ISBN: 978-91-7601-682-4 ISSN: 0346-6612 New Series No: 1889 Electronic version available at http://umu.diva-portal.org/ Printed by: UmU Print Service, Umeå University Umeå, Sweden 2017 To Sara, Viggo & Sofie

Table of Contents

Abbreviations iii Abstract iv Summary in Swedish v Original papers vii 1. Introduction 1 1.1 Microscopy 2 1.2 Molecular detection methods 3 1.2.1 Nucleic acid extraction 3 1.2.2 PCR 3 1.2.3 qPCR 4 1.2.4 PCR-based methods in diagnostic parasitology 5 1.3 Molecular methods for genetic subtyping 5 1.3.1 PCR-RFLP and riboprinting 5 1.3.2 RAPD 6 1.3.3 Sequencing 6 1.3.4 MLST 7 1.4 Metagenomics and the intestinal microbiota 7 1.5 Entamoeba histolytica and Entamoeba dispar 8 1.6 Giardia intestinalis 9 1.7 Cryptosporidium spp. 10 1.8 Dientamoeba fragilis 11 1.9 Blastocystis sp. 12 1.9.1 Taxonomy 12 1.9.2 Genetic diversity 12 1.9.3 Life cycle and transmission 13 1.9.4 Host specificity of Blastocystis subtypes 13 1.9.5 The geographic distribution of Blastocystis subtypes 15 1.9.5.1 North and South America 15 1.9.5.2 Europe 16 1.9.5.3 Africa 17 1.9.5.4 West and South Asia 18 1.9.5.5 East and Southeast Asia 19 1.9.5.6 Australia 20 1.9.5.7 The world 20 1.9.6 Pathogenicity 21 1.9.7 Microscopy 22 1.9.8 Molecular detection methods 22 1.9.9 Treatment 22

i 2. Aims 24 3. Methodological considerations 25 3.1 Paper I 25 3.2 Paper II 27 3.3 Paper III 28 3.4 Paper IV 29 4. Results and discussion 31 4.1 The sensitivity of molecular detection methods 31 4.2 PCR inhibition 32 4.3 Guanidinium thiocyanate as a transport medium 32 4.4 Blastocystis prevalence in Sweden and Zanzibar 33 4.5 Blastocystis subtypes in Sweden and Zanzibar 34 4.6 Giardia intestinalis in Zanzibar 35 4.7 Dientamoeba fragilis in Zanzibar 35 4.8 The debated pathogenicity of Blastocystis 36 4.9 Age and gender distributions of Blastocystis 36 4.10 Temporal stability of Blastocystis carriage 37 4.11 Blastocystis and the bacterial microbiota 37 4.12 Blastocystis ST4 39 5. Conclusions 41 6. Acknowledgements 42 7. References 44

ii Abbreviations

BLAST Basic local alignment search tool Ct Cycle threshold ddNTP Dideoxynucleoside triphosphate DNA Deoxyribonucleic acid dNTP Deoxynucleoside triphosphate ESBL Extended spectrum beta-lactamase FECT Formol-ether-concentration technique FDR False discovery rate Gp60 60 kDa glycoprotein Gua-SCN Guanidinium thiocyanate IBD Inflammatory bowel disease IBS Irritable bowel syndrome IgA Immunoglobulin A IPC Internal positive control LOD Limit of detection MLST Multi locus sequence typing O&P Ova and parasite PCR Polymerase chain reaction PCR-RFLP PCR restriction fragment length polymorphism PBS Phosphate buffered saline qPCR Real-time PCR RAPD Random amplification of polymorphic DNA RNA Ribonucleic acid SAF Sodium acetate-acetic acid-formalin SAR Stramenopiles, Alveolata, Rhizaria SCFA Short-chain fatty acids SSU-rDNA Small subunit ribosomal RNA gene ST Blastocystis subtype STS Sequence tagged site TFT Triple feces test TMP-SMX Trimethoprim-sulfamethoxazole WHO World Health Organization

iii Abstract

The development of molecular tools for detection and typing of unicellular intestinal parasites has revealed genetic diversities in species that were previously considered as distinct entities. Of great importance is the genetic distinction found between the pathogenic Entamoeba histolytica and the non-pathogenic Entamoeba dispar, two morphologically indistinguishable species. Blastocystis sp. is a ubiquitous intestinal parasite with unsettled pathogenicity. Molecular studies of Blastocystis sp. have identified 17 genetic subtypes, named ST1-17. Genetically, these subtypes could be considered as different species, but it is largely unknown what phenotypic or pathogenic differences exist between them. This thesis explores molecular methods for detection and genetic subtyping of unicellular intestinal parasites, with special focus on Blastocystis. We found that PCR-based methods were highly sensitive for detection of unicellular intestinal parasites, but could be partially or completely inhibited by substances present in faeces. A sample transport medium containing guanidinium thiocyanate was shown to limit the occurrence of PCR inhibition. The prevalence of Blastocystis in Swedish university students was over 40%, which is markedly higher than what was previously estimated. Blastocystis ST3 and ST4 were the two most commonly found Blastocystis subtypes in Sweden, which is similar to results from other European countries. Blastocystis sp. and Giardia intestinalis were both commonly detected in Zanzibar, Tanzania, each with a prevalence exceeding 50%. Blastocystis ST1, ST2, and ST3 were common, but ST4 was absent. While G. intestinalis was most common in the ages 2-5 years, the prevalence of Blastocystis increased with increasing age, at least up to young adulthood. We found no statistical association between diarrhoea and Blastocystis sp., specific Blastocystis subtype or G. intestinalis. Metagenomic sequencing of faecal samples from Swedes revealed that Blastocystis was associated with high intestinal bacterial genus richness, possibly signifying gastrointestinal health. Blastocystis was also positively associated with the bacterial genera Sporolactobacillus and Candidatus Carsonella, and negatively associated with the genus Bacteroides. Blastocystis ST4 was shown to have limited intra-subtype genetic diversity and limited geographic spread. ST4 was also found to be the major driver behind the positive association between Blastocystis and bacterial genus richness and the negative association with Bacteroides.

iv Summary in Swedish

Encelliga tarmparasiter som Entamoeba histolytica och Giardia intestinalis är viktiga orsaker till diarré över hela världen men framförallt i tropiska och subtropiska områden. Diagnostik av tarmparasiter har traditionellt utförts genom mikroskopi av avföringsprover. De senaste 20 åren har molekylära metoder som påvisar parasiternas arvsmassa i avföringsprov utvecklats och används allt mer för diagnostik och genetisk typning. Denna utveckling har gett upphov till flera upptäckter som föranlett omvärdering av tidigare etablerade fakta. Ett viktigt exempel är särskiljningen mellan arterna Entamoeba histolytica och Entamoeba dispar. Dessa två arter kan inte skiljas från varandra vid mikroskopi, men medan E. histolytica kan orsaka diarré, tjocktarmsinflammation och allvarliga infektioner i levern saknar E. dispar sjukdomsframkallande egenskaper. De två arterna kan dock skiljas från varandra med molekylära metoder som påvisar skillnader i arternas gener. Blastocystis sp. är en tarmparasit som är vanligt förekommande över hela världen men dess förmåga att orsaka sjukdom är ifrågasatt. Genetisk kartläggning av olika Blastocystis-stammar har identifierat 17 olika undergrupper, s.k. subtyper, som namngivits ST1–ST17. De genetiska skillnaderna mellan subtyperna är så pass stora att de skulle kunna anses vara olika arter. Det är dock inte klarlagt om subtyperna har olika egenskaper eller olika sjukdomsframkallande förmåga. Denna avhandling utforskar olika molekylära metoder för påvisning av encelliga tarmparasiter och har ett särskilt fokus på Blastocystis och dess subtyper. I studierna som ingår i denna avhandling fann vi att molekylära metoder hade betydligt högre känslighet än mikroskopi för diagnostik av Entamoeba histolytica, Giardia intestinalis och Blastocystis sp. (studie I och III). Avföringsprover innehåller dock en mängd olika ämnen som kan störa de kemiska reaktioner som används vid molekylär identifiering. Detta fenomen kallas inhibition och kan i värsta fall ge upphov till negativa analysresultat trots att arvsmassan som eftersöks finns i provet. Vi fann att inhibition var relativt vanligt förekommande och att utspädning av provmaterialet innan analys var ett enkelt sätt att undvika denna problematik. Vi utvärderade även ett provtagningsrör innehållande ämnet guanidiniumthiocyanat och fann att även detta minskade risken för inhibition. Tidigare studier utförda med mikroskopi pekade på att 5–10 % av Sveriges befolkning bar på tarmparasiten Blastocystis. Då vi undersökte avförings- prover från svenska universitetsstudenter med molekylära metoder fann vi Blastocystis hos över 40 % av studiedeltagarna (studie IV). Det är därför troligt att den allmänna förekomsten av Blastocystis i Sverige är betydligt högre än vad som tidigare ansetts. Förekomsten av Blastocystis-subtyper i Sverige var tidigare okänd och undersöktes i två studier (studie II och IV). Vi

v fann att ST3 var den vanligast förekommande subtypen och ST4 den näst vanligaste, följt av ST1, ST2 och ST7. Liknande resultat har rapporterats från andra europeiska länder. Vi använde också molekylära metoder för att påvisa encelliga tarmparasiter hos människor boende i Zanzibar, Tanzania (studie III). De två vanligaste arterna var Blastocystis sp. och Giardia intestinalis, som var och en påvisades i över 50 % av studiedeltagarna. Blastocystis ST1, ST2 och ST3 var alla vanligt förekommande medan ST4 saknades helt. G. intestinalis var vanligast bland barn i åldrarna 2–5 år medan förekomsten av Blastocystis ökade med stigande ålder, åtminstone upp till tidig vuxenålder. Vi jämförde förekomsten av tarmparasiter hos personer med och utan diarré, men fann ingen association mellan diarré och bärarskap av Blastocystis, specifik Blastocystis-subtyp eller G. intestinalis. Det är dock sannolikt att den höga förekomsten av Giardia utgör ett påtagligt hälsoproblem bland barn i Zanzibar då denna parasit kan orsaka undernäring och tillväxt- hämning även i frånvaro av diarré. I studie IV undersökte vi förhållandet mellan Blastocystis och bakteriefloran i tarmen hos svenska universitetsstudenter som reste utomlands för studier. Detta utfördes genom s.k. metagenomsekvensering som är en avancerad metod som kartlägger hela det genetiska materialet i ett prov, i det här fallet i avföringsprov. Vi fann att flera studiedeltagare hade samma Blastocystis-subtyp före och efter resa, trots utlandsvistelser som varade mellan 3 veckor och 5 månader. Detta tyder på att bärarskap med Blastocystis kan vara långvarigt. Vidare fann vi att personer med Blastocystis i genomsnitt hade ett högre antal bakteriesläkten i sin tarmflora än personer utan Blastocystis. En stor mångfald av bakteriesläkten i tarmen anses ofta vara ett tecken på god hälsa i mag-tarmkanalen och det är möjligt att Blastocystis kan vara en del av en hälsosam tarmflora. Vi fann även att Blastocystis var associerat med två specifika bakteriesläkten, Sporolactobacillus och Candidatus Carsonella, men det är oklart vilken roll dessa bakteriesläkten har i tarmen. Bärare av Blastocystis hade en lägre förekomst av Bacteroides som är ett bakteriesläkte som frodas av en kost med en hög andel animaliskt fett och protein. Detta kan antyda att det finns en koppling mellan Blastocystis och en kost rik på vegetabilier. I detta avhandlingsarbete fann vi att Blastocystis ST4 hade flera unika egenskaper. Resultaten av den genetiska typningen av ST4 antyder att ST4 i jämförelse med andra subtyper kan ha utvecklats senare i evolutionärt perspektiv. Genom att kombinera våra resultat med resultat presenterade av andra forskare kunde vi också se att ST4 är vanlig i Europa men ovanlig i Sydamerika, Afrika och Asien. I studie IV såg vi att ST4 var den subtyp som i stor grad låg bakom Blastocystis kopplingar till högt antal bakteriesläkten i tarmen och låg förekomst av Bacteroides.

vi Original papers

This thesis is based on the following papers, which will be referred to in the text by the corresponding Roman numerals.

I. Forsell J, Koskiniemi S, Hedberg I, Edebro H, Evengård B, Granlund M. Evaluation of factors affecting real-time PCR performance for diagnosis of Entamoeba histolytica and Entamoeba dispar in clinical stool samples. J Med Microbiol. 2015;64:1053-62.

II. Forsell J, Granlund M, Stensvold CR, Clark CG, Evengård B. Subtype analysis of Blastocystis isolates in Swedish patients. Eur J Clin Microbiol Infect Dis. 2012;31:1689-96. Erratum in Eur J Clin Microbiol Infect Dis. 2012;31:1697. Clark, G C [corrected to Clark, C G].

III. Forsell J, Granlund M, Samuelsson L, Koskiniemi S, Edebro H, Evengård B. High occurrence of Blastocystis sp. subtypes 1-3 and Giardia intestinalis assemblage B among patients in Zanzibar, Tanzania. Parasit Vectors. 2016;9:370.

IV. Forsell J, Bengtsson-Palme J, Angelin M, Johansson A, Evengård B, Granlund M. The relation between Blastocystis and the intestinal microbiota in Swedish travellers. Manuscript.

Reprints were made with the kind permission of the publishers.

vii viii 1. Introduction

Intestinal parasites have accompanied humans for thousands of years. Records of human infections with intestinal worms date back to ancient Greece and Rome in the period 400 BC to 200 AD [1]. The history of detection of unicellular parasites starts in 1681 when Antonie van Leeuwenhoek observed Giardia cells in his own diarrhoeal stool using a single lens microscope of his own construction [2]. In 1875, Fyodor Lesh reported his discovery of Entamoeba histolytica and its potential to cause diarrhoea in experimentally infected dogs [3]. Among the unicellular intestinal parasites there are species that are firmly implicated in human disease such as Entamoeba histolytica, Giardia intestinalis, and Cryptosporidium spp. Other species are considered non- pathogenic, e.g. Entamoeba coli, Endolimax nana, and Iodamoeba butschlii. There are also species with debated pathogenicity, such as Blastocystis sp. and Dientamoeba fragilis. Most unicellular intestinal parasites are encountered worldwide but they are more common in low-resource regions with tropical or subtropical climate and a low grade of sanitation. They are most often spread through faecal-oral transmission by ingestion of contaminated food or water. The life cycle is typically simple with a hardy cyst stage that can survive in the environment and a trophozoite stage that is active in the intestine. Although traditionally diagnosed by microscopy, the last two decades’ development of molecular tools for detection and typing of intestinal parasites has led to several advances. Genetic diversities have been revealed in genera and species previously considered as distinct entities, prompting re-evaluation of previously known facts. This thesis investigates some of these molecular methods in surveys of intestinal parasites. To give a background to the discussion of the results obtained in this thesis the introduction presents the basic concepts of microscopy and molecular methods for detection and typing of intestinal parasites, provides a brief description of the parasite species discussed: Entamoeba histolytica, Entamoeba dispar, Giardia intestinalis, Cryptosporidium spp., and Dientamoeba fragilis, followed by a more extensive description of Blastocystis sp. and its genetic subtypes – the main focus of this thesis.

1 1.1 Microscopy

Light microscopy has been the mainstay of parasite detection in faecal samples for many years. A large array of intestinal worm ova and unicellular cysts can be detected in a diagnostic test that is often referred to as the ova and parasite (O&P) exam. Faecal samples are most often collected in a fixative, such as sodium acetate-acetic acid-formalin (SAF), to preserve the morphology of cysts and trophozoites [4,5]. Fresh samples collected without fixatives can also be examined to detect motility in trophozoites, but this is often omitted in routine diagnostics. Slides can be prepared directly from faeces, but the detection of cysts is facilitated by concentration of the faecal samples prior to microscopy, for instance by the formol-ether-concentration technique (FECT) that separates the parasites from the fat and debris present in faeces [6]. Cysts can be detected in wet mounts, either unstained or stained with an iodine solution (Fig. 1). The use of permanent stains such as trichrome [7], iron haematoxylin [8], or chlorazol black [9, 10] allows for the detection of both trophozoites and cysts under oil immersion high power magnification (Fig. 1). Acid fast staining, such as Ziehl-Neelsen staining [11, 12], facilitates the detection of coccidian oocysts of genera Cryptosporidium, Cyclospora and Cystoisospora. Since the shedding of parasites in faeces can be intermittent, sampling on three separate occasions is usually recommended to improve detection sensitivity. The sensitivity of investigating one individual stool sample compared to examination of three stool samples has been shown to be around 75% [13, 14]. Van Gool et al [10] described a “triple feces test” (TFT), which included sampling on three occasions, using two tubes with SAF and one tube without preservatives, and microscopic detection using both iodine staining and permanent staining with chlorazol black. The detection sensitivity of the TFT was at least twice as high as microscopy of one unfixed faecal sample only stained with iodine.

Fig. 1. Blastocystis sp. in an iodine stained wet smear (left) and a permanently stained Giardia intestinalis trophozoite under high power magnification (right)

2 1.2 Molecular detection methods

Molecular detection methods detect nucleic acid, i.e. DNA or RNA. These methods are of great value in clinical microbiology for the detection of bacteria, viruses, fungi and parasites.

1.2.1 Nucleic acid extraction

Molecular detection in clinical samples is preceded by methods that extract the nucleic acid from the other molecules present in the organism and sample. Traditional methods include the phenol-chloroform method [15] and the method developed by Boom et al [16] using guanidinium thiocyanate and silica particles. There are also a multitude of commercial methods available, including manual methods and automated systems. The major components of nucleic acid extraction are lysis of cells, inactivation of cellular nucleases such as DNAase and RNAase, and separation of the nucleic acid from the other molecules [17]. Of importance in the field of intestinal parasites is the ability of the extraction method to break down the cyst wall to enable the release of the nucleic acid.

1.2.2 PCR

The prototypical molecular detection method is the polymerase chain reaction, or PCR. The method was developed in the 1980’s, and is credited to Kary Mullis [18] who later received the Nobel Prize in chemistry for his invention. PCR allows multiplication of a specific DNA sequence by a factor of 109-1012 in less than two hours. The reaction is typically carried out in small tubes which are placed in a thermocycler that has the ability to rapidly change temperature. The reaction requires template DNA, a thermostable DNA polymerase, two primers, deoxynucleoside triphosphates (dNTPs), and Mg2+, all added to a buffer. The two primers are short nucleotide sequences that are complementary to the 3' ends of the targeted sequence on each of the two DNA strands. The dNTPs (dATP, dGTP, dCTP, and dTTP) are the building blocks required to synthesize new DNA. The amplification of target DNA is achieved by the repeated cycling of three steps – denaturation, annealing, and extension. Denaturation occurs at around 95oC, cleaving the double stranded DNA into two single strands. Annealing takes place at 50-65oC in which the primers bind to their complementary sites of the single strands, acting as a starting point for the DNA polymerase. Extension is typically achieved at 72oC where the DNA polymerase synthesizes a new DNA strand, complementary to the template

3 strand, by successively adding dNTPs. The process is then repeated for 30- 40 cycles with each cycle doubling the amount of target DNA from the previous cycle. The result of the amplification, the PCR product, is then subjected to gel electrophoresis, and visualised by staining, to determine its size. The assay is considered positive for the DNA target if the PCR product is of the expected size. The specificity of the assay relies on the design of the primers and their binding to the intended DNA sequence.

1.2.3 qPCR

Real-time PCR (qPCR) is a development of the PCR [19] performed in specialised instruments in which amplification of target DNA is measured in real-time and presented by a computer software. This can be achieved by detecting the emitted fluorescence of dyes that bind to double stranded DNA. Further specificity is obtained by using molecular probes specific to the DNA target. Several probe technologies exist, and a common variant is the Taqman probe [20] which is used in the qPCR assays described in this thesis. The Taqman probe is an oligonucleotide that is complementary to a specific sequence of the DNA target, and has a fluorophore and a quencher molecule bound to it. The fluorophore emits no fluorescence as long as it is in close proximity to the quencher. In the annealing phase of the PCR, the probe also binds to the DNA target along with the primers. In the extension phase, the DNA polymerase destroys the bound probe as it synthesizes new DNA. This leads to a separation of the fluorophore and quencher and the emitted fluorescence can be detected. Duplex or multiplex assays for simultaneous detection of two or more targets in one reaction are common. This is achieved by adding primers and probes for all the targets in the same reaction mixture, with each probe having a unique fluorophore, allowing for specific target detection. qPCR simplifies the workflow since no handling of the PCR product for visualisation is needed. This also limits the possibility of laboratory contamination with amplified PCR products. The detected fluorescence is displayed as an amplification curve in which the cycle threshold (Ct) is the cycle in which a positive reaction clearly separates from the background fluorescence. The Ct value can be compared to a DNA standard with known concentrations of the target gene to quantify the amount of target present at the start of the reaction. Even without a DNA standard to compare with, the Ct value is an indicator of the amount of target DNA in the sample, where lower Ct values indicate higher amounts of target and higher Ct values indicate lower amounts.

4 1.2.4 PCR-based methods in diagnostic parasitology

Several in-house and commercial PCR and qPCR assays have been described for the unicellular intestinal parasites that are included in this thesis, as reviewed by Verweij and Stensvold [21]. The major advantage of PCR-based methods is the high sensitivity and specificity achieved in comparison with microscopy [21, 22]. The major drawback with molecular detection methods are that they only detect the organisms that they have been designed to detect. Microscopy on the other hand offers the possibility to detect a multitude of different species of parasites, including very rare organisms. Another potential drawback with performing PCR on faecal samples is that inhibitory substances present in faeces can be co extracted with the DNA and cause false negative results. The need for specialised equipment limits the use of PCR diagnostics in low resource settings in which intestinal parasites are most common. Instrument and reagent costs can be high for qPCR diagnostics but the hands-on time compared to microscopy is rather low, making PCR-based diagnostics attractive in settings with high labour costs. During the last decade, qPCR assays to detect intestinal parasites have increasingly been implemented in diagnostic laboratories in Europe. This has led to a markedly increased detection of Giardia intestinalis and Cryptosporidium [22], offering a diagnosis in cases of diarrhoea that would otherwise be unresolved. In the modern diagnostic laboratory, microscopy and molecular detection methods complement each other well, offering broad detection range and high sensitivity respectively.

1.3 Molecular methods for genetic subtyping

Different methods have been developed for the purpose of detecting genetic differences between organisms or between different strains of the same organism. The methods that are important in the context of this thesis are presented below.

1.3.1 PCR-RFLP and riboprinting

PCR restriction fragment length polymorphism (PCR-RFLP) uses restriction enzymes [23] to further characterise PCR products. The restriction enzymes cut DNA at specific nucleotide sequences known as restriction sites. Different DNA sequences will have different number and location of the restriction sites, resulting in varying number and sizes of DNA fragments. The fragments are separated according to size by gel electrophoresis which

5 creates an RFLP-pattern that can be compared to patterns of homologous PCR products. PCR-RFLP specifically targeting ribosomal genes is sometimes referred to as riboprinting [24].

1.3.2 RAPD

Randomly amplified polymorphic DNA (RAPD) is a PCR performed with several arbitrary primers that amplify targets of unknown locations of a genomic DNA template [25]. The products are then separated by gel electrophoresis for pattern analysis.

1.3.3 Sequencing

DNA sequencing is used to determine the nucleotide bases and their precise order in a sequence. This offers the possibility to find discrete differences between homologous genes. A commonly used method is dye-terminator sequencing [26]. This method comprises a PCR, with DNA polymerase, primers, and deoxynucleoside triphosphates (dNTPs), with the addition of modified dideoxynucleoside triphosphates (ddNTPs). Each of the four ddNTPs is labeled with a unique fluorescent dye and they all terminate polymerization once they are incorporated into the synthesized strand. The concentration of ddNTPs is lower than that of dNTPs, and they are incorporated at random. This results in a mix of synthesized sequence fragments of varying sizes with a labeled nucleotide at the end. The mix of DNA fragments are then passed through a capillary tube in which the smallest fragment, representing the beginning of the sequence, comes first and the largest last. Fluorescence is emitted by laser excitation of the molecules as they pass, and this fluorescence is detected by a detector system, creating a chromatogram that describes the nucleotide sequence (Fig. 2.). A common target in DNA sequencing is the complete or partial sequence of the small subunit unit rRNA gene (SSU-rDNA).

Fig. 2. An example of a sequencing chromatogram.

6 1.3.4 MLST

Multi locus sequence typing (MLST) has been widely used to detect genetic differences within species to investigate population structures and compare strains, especially for bacteria [27], but also for parasites [28]. In MLST, several genes or gene fragments are sequenced for each strain and each sequence is assigned to an allele. The combined allelic profile of all the sequences of a strain determines the MLST sequence type.

1.4 Metagenomics and the intestinal microbiota

The metagenome is the collective genetic material from all organisms present in a sample [29, 30]. Data for metagenomic analysis is obtained by advanced high throughput sequencing, in which a massive amount of relatively short nucleotide sequences are obtained. These sequences can be assigned to genes by computational analyses. Comparisons of the metagenomic data to known sequences of SSU-rDNA (16S-rDNA for prokaryotes and 18S-rDNA for eukaryotes) reveal which genera or species are present in a sample, and their relative abundance. Metagenomic sequencing of faecal samples can be used to assess the composition of all the microorganisms that are present in the intestine, i.e. the intestinal microbiota. This microbiota is dominated by bacteria belonging to the divisions Bacteroidetes, Firmicutes, and Actinobacteria [30]. Archaea and eukaryotes are present in smaller amounts [31]. Several factors can influence the composition of the intestinal microbiota such as diet, age and antibiotic treatment [32]. Although approximately 400 bacterial species have been cultured from the human intestine, metagenomic analyses have revealed the presence of over 1000 phylotypes (species equivalents) [33]. The sheer number of species as well as inter- and intra-individual differences make interpretations of metagenomic data from the intestine difficult. In 2011, Arumugam et al [34] reported that the human intestinal microbiota could be stratified into three major enterotypes, each signified by an enrichment of either Bacteroides, Prevotella or Ruminococcus. The existence of enterotypes enriched in Bacteroides or Prevotella have since been validated by other researches and a meta-analysis of studies on the intestinal microbiota confirms an enrichment of Bacteroides in the microbiota of people living in Western societies and an enrichment of Prevotella in people living in agricultural low-resource regions [35]. Many diseases have been linked to alterations in the intestinal microbiota, including diabetes [36, 37], cardiovascular disease [38], obesity [39, 40], inflammatory bowel disease (IBD) [41, 42, 43] and irritable bowel syndrome

7 (IBS) [44]. The underlying mechanisms for these associations are presently being investigated by several research groups. Microbial production of short- chain fatty acids (SCFAs), such as butyrate and acetate from dietary starch, is thought to be among the substances that can affect the immunometabolic functions of the host [45, 46].

1.5 Entamoeba histolytica and Entamoeba dispar

Entamoeba histolytica belongs to the phylum Amoebozoa [47], and as the species name implies is an intestinal amoeba that can cause tissue destruction. E. histolytica occurs worldwide, but it is more common in areas with poor sanitation, particularly in the tropic regions of South and Central America, Africa, and Asia. Entamoeba histolytica can cause diarrhoea, severe amoebic colitis, and extra-intestinal disease, primarily amoebic liver abscess. It is responsible for considerable morbidity and mortality worldwide, causing up to 100.000 deaths each year [48]. The life cycle is simple, with a hardy cyst stage that can survive in the environment and contaminate water or food, and a trophozoite stage that is active in the intestine, causing symptoms. Humans are the only know hosts of the parasite and it can therefore be eradicated from a community by rigorous sanitation. In non-endemic countries risk groups for infection are travellers, immigrants from endemic regions, and men who have sex with men [49]. A high occurrence of asymptomatic carriage with E. histolytica had been observed for many decades before the parasite was formally redescribed in 1993 as two separate species, the pathogenic E. histolytica and the non- pathogenic Entamoeba dispar [50]. The cysts of these two species are morphologically indistinguishable from each other and 80-90% of microscopy positive cases are in fact E. dispar [51]. Discrimination of the two species is recommended by the World Health Organization (WHO) to avoid unnecessary treatment of E. dispar [48]. This can be accomplished by PCR-based methods if the resources are available. Although asymptomatic carriage of E. dispar does not require treatment, asymptomatic carriage of E. histolytica does require treatment to prevent later development of invasive disease. A luminal agent, either paromomycin, diloxanide furoate, or iodoquinol, is recommended for this purpose. Invasive disease is treated with metronidazole followed by a luminal agent [52].

8 1.6 Giardia intestinalis

Giardia intestinalis, sometimes referred to as Giardia lamblia or Giardia duodenalis, is a flagellate belonging to the phylum Metamonada [47]. It has a worldwide distribution, but is more common in tropic and subtropic regions with low levels of sanitation. In regions of endemicity it is especially common among children, where it is an important cause of diarrhoea and malabsorption. The World Health Organization (WHO) has therefore included Giardia in the ‘Neglected Diseases Initiative’ [53] which aims to improve health and socio-economic development in affected regions. The parasite has a simple cyst and trophozoite life cycle. Giardia intestinalis can be found in a large variety of animals which can contaminate the environment with cysts, potentially causing zoonotic transmission. In Sweden, around 1500 Giardia-infections are reported each year (15/100.000 inhabitants), of which approximately 200 are considered domestically transmitted [54]. Giardia-contamination of water supplies can occasionally cause large outbreaks of diarrhoeal disease infecting thousands of individuals, such as those affecting Sälen, Sweden in 1986 [55], and Bergen, Norway in 2004 [56]. Smaller outbreaks can occur in the context of outdoor recreational activities, by consumption of contaminated food, and in day- care centres [57, 58]. Studies of the genes encoding β-giardin, triose phosphate isomerase, glutamate dehydrogenase and SSU-rRNA in G. intestinalis isolates have identified seven major genetic subgroups, named assemblage A to G [59]. Almost all human Giardia infections are caused by assemblage A or B [60, 61]. These assemblages also have a large number of identified animal hosts, whereas assemblages C-H seem to have a narrower animal host range [60]. The genomes of assemblage A and B share only 77% identity in protein coding regions, which suggest that they might represent two separate species [62]. In humans, some studies associate assemblage A with diarrhoea [63, 64], while others associate assemblage B with diarrhoea [65, 66]. Treatment choices for Giardia-infections include tinidazole as a single dose or metronidazole for 5-7 days [52]. Asymptomatic carriers in non- endemic areas should be treated to limit transmission of the parasite and to avoid chronic infection and malabsorption. Treatment of asymptomatic carriers in areas of high endemicity is complex, considering the frequency of reinfections on one side and the detrimental effects caused by malabsorption on the other.

9 1.7 Cryptosporidium spp.

Cryptosporidium spp. are coccidian parasites of the phylum Apicomplexa [47] that have a worldwide distribution. Humans are primarily infected with the species C. hominis and C. parvum, but twelve other species have also been detected in humans, including C. cuniculus. C. felis, C. meleagridis, and C. viatorum [67]. First primarily considered as an important pathogen in livestock, human infections received attention in the 1980s when Cryptosporidium was seen causing prolonged, severe, and sometimes fatal diarrhoea in AIDS-patients. It has since been recognised as a cause of watery diarrhoea also in immunocompetent individuals, where the disease is self-limiting. In low- income regions, infections primarily occur in small children, causing considerable morbidity and mortality due to diarrhoea [68], and cryptosporidiosis is also included in WHO’s ‘Neglected Diseases Initiative’ [53]. The life cycle of Cryptosporidium is relatively complex. The oocyst is the infective stage that can survive in the environment and is resistant to chlorine at concentrations used for water treatment. After ingestion, the oocyst releases sporozoites that invade intestinal epithelial cells, where they mature and undergo both asexual and sexual reproduction. Sexual reproduction leads to the development of new oocysts that sporulate. Some of these oocysts release their sporozoites within the host causing autoinfection, while other oocysts are shed in faeces and are directly infective to other hosts [69]. The transmission routes of Cryptosporidium – water-borne, food-borne, zoonotic, and person-to-person – are the same as those of Giardia intestinalis. The world’s largest water-borne outbreak of cryptosporidiosis occurred in 1993 in Milwaukee, USA, causing diarrhoea in 400.000 individuals [70]. Recent years has seen two large outbreaks in Sweden, in 2010 in Östersund [71], and in 2011 in Skellefteå [72], that together affected around 47.000 individuals. This can be compared to the around 500 Cryptosporidium-cases that are normally reported in Sweden each year [54]. The different species of Cryptosporidium are indistinguishable from each other by microscopy. Oocysts of Cryptosporidium spp. are half the size of other parasitic cysts and the microscopic detection of them in unstained or iodine-stained preparations are dependent on the skill and training of the examiner. Acid fast-staining facilitates diagnosis and is often used when Cryptosporidium is suspected. Cryptosporidium species determination can be achieved by PCR-RFLP or sequencing of SSU-rDNA [73]. A common method for genetic subtyping of C.

10 hominis and C. parvum is sequencing of the 60 kDa glycoprotein (Gp60) gene [74, 75]. Few antimicrobials have shown effect on Cryptosporidium, and specific treatment is usually only considered in severe cases. Nitazoxanide has been shown to shorten the duration of diarrhoea caused by Cryptosporidium in immunocompetent individuals but not in patients with HIV that are not under effective antiretroviral therapy [76, 77].

1.8 Dientamoeba fragilis

Dientamoeba fragilis, an unflagellated member of the phylum Metamonada, is an intestinal parasite predominately found in humans. Its role in disease is debated, but it has been associated with gastrointestinal symptoms such as diarrhoea, nausea, and abdominal pain [78], especially among children [79, 80, 81]. However, asymptomatic carriage is common in all age groups [82, 83]. Most research on Dientamoeba has been conducted in high-income countries, where the parasite, in contrast to most other intestinal parasites, also seems to have the highest prevalence [84]. A peculiarity with D. fragilis is the apparent lack of a cyst stage. Since the trophozoite stage is fragile and degenerates after shedding in faeces there are unanswered questions regarding the life cycle and transmission of the parasite. One theory is that the trophozoites are co-transmitted with the eggs of the intestinal worm Enterobius vermicularis, and DNA of D. fragilis has in fact been detected in Enterobius eggs [85, 86]. A recent microscopy study has detected Dientamoeba cysts in human faeces, which were half the size of the trophozoites and had a thick cyst wall [87]. These cysts were very rare and it is not known if they have a role in transmission. Since the cyst stage is absent or very rare, diagnostic microscopy for D. fragilis relies on the detection of trophozoites. Faecal samples need to be collected in a fixative, e g SAF, as to limit the destruction of the trophozoites during transport. The trophozoites are not readily seen in wet smears so a permanent staining with iron-haematoxylin, trichrome, or chlorazol black [10, 88] is needed. Examination of permanently stained smears is rather time consuming and PCR-based detection is a possible alternative [84]. There appears to be little genetic diversity in D. fragilis, with two genotypes recognised based on differences in the SSU-rDNA, named genotype 1 and genotype 2 [89]. Treatment of Dientamoeba can be considered in symptomatic patients were all other causes of gastrointestinal disease have been ruled out. The luminal agents iodoquinol and paromomycin have shown rather good cure rates, and the efficacy of metronidazole is often high, but there are reports of treatment failures with this drug [90].

11 1.9 Blastocystis sp.

First described by Alexeieff in 1911 [91], Blastocystis is the most commonly found non-fungal eukaryotic organism in human faecal samples [92, 93]. Even though it is well known and common, many aspects of Blastocystis are not fully understood. Most importantly, its ability to cause human disease is a subject for a long and unsettled debate.

1.9.1 Taxonomy

Blastocystis sp. is an anaerobic unicellular eukaryote that belongs to the phylum Stramenopiles [94] of the SAR (Stramenopiles, Alveolata, Rhizaria) supergroup [47]. Stramenopiles is a heterogeneous phylum which contains both unicellular and multicellular organisms, including many water-living organisms such as diatoms and brown algae. Blastocystis is the only member of Stramenopiles that has been found in the human intestine. In fact, only one other member of the Stramenopiles have been found infecting humans, Pythidium insidiosum, the cause of the rare and deadly tropical disease pythiosis [95]. The species name previously used for all Blastocystis found in humans was Blastocystis hominis. Since the realisation that genetic subtypes of Blastocystis can be shared by humans and animals, all findings of the parasite are now denominated Blastocystis sp., irrespective of host [96].

1.9.2 Genetic diversity

In 1997, Clark reported extensive genetic diversity in 30 human Blastocystis isolates as determined by riboprinting, identifying 7 distinct patterns, or ribodemes, using 12 restriction enzymes [97]. The same year, Böhm-Gloning et al [98] reported five distinct subgroups in 166 human Blastocystis isolates that had been subjected to riboprinting with 3 restriction enzymes. Genetic diversity was also observed by Yoshikawa et al [99] using random amplified polymorphic DNA (RAPD). Further studies of the genetic diversity using riboprinting [100, 101, 102] and the primers derived from the RAPD [100, 102] were followed by studies that performed sequencing of the SSU-rDNA [103, 104, 105], all confirming the genetic diversity in Blastocystis. In 2006, Scicluna et al [106] described the ‘DNA barcoding’ method for Blastocystis, in which ~600 bp of the SSU-rDNA is sequenced, as an efficient tool for Blastocystis subtyping. Owing to differences in methodologies the terminology differed between studies and was not easily translatable. A consensus was achieved in 2007 in which the previously reported ribodemes, subtypes, subgroups, clusters and

12 clades of Blastocystis were converted into nine distinct subtypes, named ST1-9 [96]. Since then, novel subtypes, ST10-17, have been detected in various animal species [107, 108, 109, 110]. The consensus terminology of subtypes has been used throughout this thesis, and results from older studies have been translated accordingly. The high level of genetic diversity found between Blastocystis subtypes could suggest that they in fact constitute different species [96], but it is largely unknown what phenotypic or pathogenic differences exist between them.

1.9.3 Life cycle and transmission

Microscopic examinations of Blastocystis have revealed several morphological forms of the organism, including vacuolar, granular, amoeboid, and cyst form [111]. What functions these forms have is largely unknown, and a number of disparate life cycles have been proposed [111- 113]. However, none of these life cycles have been conclusively demonstrated. Blastocystis is presumed to be transmitted by the faecal-oral route [111]. Drinking of untreated water has been implicated as a source of transmission in studies from Thailand and China [114-116]. The role of food-borne transmission has not been widely researched, but Blastocystis has been detected in leafy vegetables in Saudi Arabia [117] and a study from China associated the consumption of raw water plants specifically to Blastocystis ST1 [116]. The occurrence of person-to-person transmission is unclear. In a study from Australia, all household contacts of 11 symptomatic Blastocystis- carriers also harboured Blastocystis [118], suggesting person-to-person transmission or a common transmission source. In contrast, an epidemiological survey of 50 households in the USA detected Blastocystis in ten individuals, but found no occurrence of two or more Blastocystis-carriers within the same family [119]. Subtyping studies have found suggestive evidence of zoonotic transmission, especially among animal handlers, but the direction of transmission between humans and animals is difficult to determine [107, 108].

1.9.4 Host specificity of Blastocystis subtypes

The prevalence of Blastocystis is often high in farm animals such as cattle, pigs, and chickens [120, 121]. This is possibly a reflection of a high frequency of faecal-oral transmission in confined areas. Occurrence of Blastocystis in mammalian carnivores seems to be low [108, 110, 122]. For dogs, complete

13 absence of Blastocystis was reported in a study from Japan [120], and a low prevalence in dogs was seen in both Cambodia (1.5 %) and Australia (2.3%) [123]. A relatively high prevalence was observed in stray dogs in India (24%), possibly due to coprophagy caused by their living conditions [123]. All Blastocystis subtypes found in humans, except the rare ST9, have also been found in various animals. In Table 1, the known mammalian and avian Blastocystis subtypes, described in study compilations by Alfellani et al [110] and Cian et al [122], are presented together with their occurrence in humans. Even though many subtypes are found both in humans and animals, and zoonotic transmission is likely to occur in certain cases, there are often intra- subtype sequence variations between hosts indicating that human carriage of Blastocystis is primarily caused by anthroponotic transmission [110]. Blasocystis ST5 and ST10 are common in pigs and cattle respectively, but rare or absent in humans, indicating that zoonotic transmission of these subtypes is very limited [110, 122].

Table 1. Blastocystis subtypes in humans and animals

Subtype Occurrence Common animal hostsb in humansa ST1 +++ Non-human primates, pigs, cattle ST2 ++ Non-human primates, pigs ST3 +++ Non-human primates ST4 ++ Rodents ST5 (+) Pigs, camels ST6 + Birds ST7 + Birds ST8 (+) Non-human primates ST9 (+) - ST10 - Cattle ST11 - Elephants, non-human primates ST12 (+) Giraffes ST13 - Non-human primates, deer ST14 - Cattle, camels ST15 - Elephants ST16 - Kangaroos ST17 - Rodents a) Blastocystis subtypes in humans are denoted by increasing occurrence from (+) to +++, based on the global subtype distribution described in section 1.9.5 of this thesis. b) Several of the subtypes, especially STs 1-8, have also been detected to a lesser extent in other animal hosts, reviewed by Alfellani et al [110] and Cian et al [122]. - indicates no detection

14 1.9.5 The geographic distribution of Blastocystis subtypes

The interest in the genetic diversity of Blastocystis has led to the inclusion of subtyping in numerous Blastocystis studies all over the world. The accumulated data has here been used to describe the human Blastocystis subtype distributions in different geographical regions. All types of studies have been included, and data emanates from both symptomatic and asymptomatic individuals.

1.9.5.1 North and South America

The Blastocystis subtype distribution in North and South America is shown in Fig. 3. Data was compiled from 1011 subtype observations presented in 14 studies, representing samples from Argentina [124, 125], Bolivia [125, 126], Brazil [125, 127, 128], Colombia [125, 129, 130], Ecuador [125, 131], Mexico [132-135], Peru [125], and the USA [119, 136]. While ST3 was the dominant subtype in most study settings, ST1 was the most commonly found subtype in the three studies from Colombia [125, 129, 130], and in one study each from Brazil [127], Ecuador [125], and Mexico [133]. In one study from Bolivia [125], ST2 was the most common, and that study also included the first human report of ST12.

Fig. 3. The Blastocystis subtypes observed in North and South America.

15 1.9.5.2 Europe

The Blastocystis subtype distribution in Europe is shown in Fig. 4. Data was compiled from 1710 subtype observations presented in 24 studies (including papers II and IV), representing samples from Denmark [137-143], France [144-146], Germany [98, 147], Greece [148], Ireland [149, 150], Italy [151, 152], the Netherlands [153], Spain [154], Sweden [paper II, paper IV], and the UK [97, 106, 155]. ST3 was the most prevalent subtype in all but five studies. These five studies originated from Denmark [141, 143], France [145], Spain [154], and Sweden [paper IV] and reported ST4 as the most commonly detected subtype.

Fig. 4. The Blastocystis subtype distribution in Europe. ST12 has not been found in this region.

16 1.9.5.3 Africa

The Blastocystis subtype distribution in Africa is shown in Fig. 5. Data was compiled from 672 subtype observations presented in 10 studies (including paper III), representing samples from Egypt [156-159], Liberia [155], Libya [155, 160], Nigeria [155, 161], Senegal [162], and Tanzania [163, paper III]. ST3 was the most commonly found subtype in Egypt, Liberia, and Senegal and ST1 was the most common subtype in Nigeria, Libya, and Tanzania.

Fig. 5. The Blastocystis subtype distribution in Africa. ST5, ST8, ST9, and ST12 have not been found in this region.

17 1.9.5.4 West and South Asia

The Blastocystis subtype distribution in West and South Asia is shown in Fig. 6. Data was compiled from 1567 subtype observations presented in 26 studies, representing samples from Bangladesh [147], India [164, 165], Iran [166-171], Lebanon [172, 173], Nepal [174, 175], Pakistan [147, 176], Qatar [177], Turkey [178-187], and the United Arab Emirates [188]. ST3 was the most prevalent subtype in most studies. ST1 was the most common subtype in one study from Iran [166], one study from Pakistan [176], and two studies from Turkey [183, 185], while ST4 was the most common subtype in another study from Iran [169]. ST6 was the most commonly detected subtype in one study from Nepal [175].

Fig. 6. The Blastocystis subtype distribution in West and South Asia. St8, ST9, and ST12 have not been found in this region.

18 1.9.5.5 East and Southeast Asia

The Blastocystis subtype distribution in East and Southeast Asia is shown in Fig. 7. Data was compiled from 1452 subtype observations presented in 21 studies, representing samples from Cambodia [189], China [116, 190, 191], Indonesia [192], Japan [100, 101, 147], Malaysia [193, 194], the Philippines [195-197], Singapore [198], and Thailand [115, 147, 199-204]. ST3 was the most prevalent subtype in most studies. ST1 was the most common subtype in three studies from Thailand [115, 199, 201], and in one study each from Indonesia [192] and the Philippines [195].

Fig. 7. The Blastocystis subtype distribution in East and Southeast Asia. ST 8 and ST12 have not been found in this region.

19 1.9.5.6 Australia

The Blastocystis subtype distribution in Australia is shown in Fig. 8. Data was compiled from 135 subtype observations presented in 3 studies [118, 189, 205].

Fig. 8. The Blastocystis subtype distribution in Australia. ST9 and ST12 have not been found in this region.

1.9.5.7 The world

Compiling the data presented above results in the following global Blastocystis subtype distribution: ST1 29.6% (n = 1967), ST2 13.8% (n = 901), ST3 43.6% (n = 2854), ST4 7.6% (n = 498), ST5 0.7% (n = 44), ST6 2.6% (n = 167), ST7 1.8% (n = 119), ST8 0.2% (n = 13), ST9 0.2% (n = 11), and ST12 0.04% (n = 3). ST3 is clearly the most commonly detected subtype, and STs 1-4 represent over 94% of all human subtype observations. The most notable geographic difference in subtype distributions is that ST4 is the second most common subtype in Europe and the third most common subtype in Australia, but only rarely found in the rest of the world.

20 1.9.6 Pathogenicity

Blastocystis has been associated to diarrhoea, flatulence, abdominal pain, and gastrointestinal discomfort [92, 93]. There are several reports of patients with Blastocystis as the only detected possible pathogen that experienced cessation of symptoms after successful treatment of the parasite [206-208]. However, Blastocystis role in human diseases is frequently debated. Many studies have reported equal Blastocystis occurrence in both symptomatic and asymptomatic individuals [209-212]. Since a high degree of genetic diversity exists between Blastocystis subtypes the debated pathogenicity of the parasite could be caused by certain subtypes being pathogenic and others being non-pathogenic. This has been the focus of studies that have compared subtype distributions in symptomatic and asymptomatic groups. ST1 has been linked to symptomatic carriage in several smaller studies [156, 172, 180, 190] but also to asymptomatic carriage [130]. Similar conflicting reports can be found for ST2 [130, 179] and ST3 [160, 180, 190]. Other studies report no significant differences in subtype distribution between symptomatic and asymptomatic carriers of Blastocystis [172, 182]. Two studies from Spain [154] and Denmark [141] report a high prevalence of ST4 in symptomatic patients, but these studies included no healthy controls. The symptoms sometimes associated with Blastocystis are similar to those of irritable bowel syndrome (IBS), and studies have investigated associations between the two. In fact, several studies have shown significantly higher prevalence of Blastocystis in IBS patients than in controls [176, 213-215]. On the other hand, there are also studies that have reported no significant differences in Blastocystis carriage between IBS patients and controls [171, 216-218]. Some studies have observed a quite clear overrepresentation of a specific Blastocystis subtype in IBS patients. However, the implicated subtypes differ between study settings: ST1 in Pakistan [176] and Egypt [158], ST3 in Colombia [130], and ST4 in Italy [152], making interpretations of these results difficult. Other studies report no significant differences in subtype distributions between IBS patients and controls [132, 171, 218]. In vitro studies have found some potential mechanisms for Blastocystis pathogenesis via the expression of cysteine proteases. These include the degradation of human secretory IgA (immunoglobulin A) [219], induction of apoptosis in intestinal epithelial cells and disruption of the epithelial barrier function [220], as well as activation of the cytokine interleukin 8 in colonic epithelial cells [221]. These experiments were performed with Blastocystis ST4 and it has been observed that ST7 has even greater protease activity [222]. ST7 proteases have subsequently also been shown to disrupt the epithelial barrier function in intestinal cells in vitro [223]. Rats experimentally infected with Blastocystis ST1 from a diarrhoeic human

21 showed variable histopathological changes of the intestine, with some rats exhibiting mucous membrane injury and parasite invasion into the mucosa or lamina propria [224]. As it stands, the pathogenicity of Blastocystis is still open for debate. There is also no clear distinction between pathogenic and non-pathogenic subtypes.

1.9.7 Microscopy

Like other intestinal parasites, the shedding of Blastocystis in stool can be intermittent and the collection of three faecal samples is recommended [10]. Because of its polymorphic nature, Blastocystis may be overlooked in microscopy [93], and microscopic detection using wet smears, either performed directly or after FECT, have low sensitivity [138, 225-226]. Examination of smears permanently stained with trichrome increases the sensitivity [138, 226,]. The detection sensitivity is also markedly increased by the use of short term xenic culture [138, 225-226].

1.9.8 Molecular detection methods

Sensitive detection of Blastocystis can be achieved by qPCR, and two different qPCR assays targeting the SSU-rDNA of Blastocystis have been described by Poirier et al in 2011 [145] and by Stensvold et al in 2012 [227]. The assay developed by Poirier et al amplifies a ~320 bp product that can be sequenced for Blastocystis subtype designation. The assay by Stensvold et al is possibly more sensitive, owing to its smaller product size of around 115 bp, but it cannot be used for subtyping.

1.9.9 Treatment

Considering the unsettled pathogenicity of Blastocystis, treatment is generally not recommended to asymptomatic carriers. Treatment can be considered in cases of prolonged gastrointestinal symptoms were Blastocystis is the only possible pathogen detected and other non-infectious causes of the symptoms have been ruled out. Treatment failures are common, and complete eradication of Blastocystis can be difficult to achieve [118, 228]. Several antimicrobials have been used in the treatment of Blastocystis, with varying degree of scientific documentation. These include metronidazole, trimethoprim-sulfamethoxazole (TMP-SMX), nitazoxanide,

22 paromomycin, secnidazole, furazolidone, iodoquinol, ketoconazole, tinidazole, and the probiotic yeast Saccharomyces boulardii [228, 229]. Metronidazole is the most commonly prescribed antimicrobial for Blastocystis. This drug has shown good effect in case reports [230, 231] and in a placebo-controlled trial, were cessation of symptoms and parasite clearance was observed in 88% and 80% respectively, both significantly higher than what was observed for placebo [232]. However, treatment failures [118, 207, 228, 233] and modest cure rates [234] have also been reported. In vitro studies have confirmed the existence of both metronidazole resistant and susceptible isolates of Blastocystis [235, 236]. Another treatment option is TMP-SMX, used either as stand-alone treatment or in combination with metronidazole. TMP-SMX has shown good treatment efficacy against Blastocystis [207, 237], but treatment failures [118] and low cure rates in high density infections [234] have also been reported. The effect of nitazoxanide has been studied in two placebo-controlled trials, in which a significantly higher cure rate over placebo was observed in one of them [238], but not in the other [239]. The luminal agent Paromomycin has shown good clinical effect in case reports [240, 241] and small studies [228, 233]. Triple therapy with secnidazole, furazolidone, and nitazoxanide has been used in certain clinics, perhaps as a last resort, but treatment failures have been observed for this treatment as well [228]. Treatment with the yeast Saccharomyces boulardii has shown similar efficacy as metronidazole in one study [242]. In the largest in vitro Blastocystis susceptibility study to date, 12 cultured Blastocystis isolates (ST1, ST3, ST4, ST8) were tested against 12 antimicrobials [243]. In the study, metronidazole and secnidazole were found to have an effect on Blastocystis but only at high concentrations, which limits the potential for eradication when these drugs are used in safe dosages. Nitazoxanide, furazolidone, paromomycin, and the antifungal drugs fluconazole, nystatin and itraconazole all had little to no effect on Blastocystis. The antihelminth drugs albendazole and ivermectin had effect at high concentrations. The best in vitro effect was found for TMP-SMX, which the authors therefore recommended as first-line therapy. Some subtype differences in drug susceptibility have been noted [235, 243], but the studies are too small to draw any firm conclusions on susceptibility differences between the subtypes. Evidently, there is room for further studies addressing susceptibility testing for the most common drugs against Blastocystis and Blastocystis subtypes.

23 2. Aims

The aims of this thesis were to evaluate molecular methods for detection of unicellular intestinal parasites, study the distribution of genetically defined subtypes of Blastocystis in areas not previously studied, and to investigate the relationship between Blastocystis and the bacterial microbiota.

The specific aims were:

To characterize factors affecting the performance of a qPCR-assay to detect Entamoeba histolytica and Entamoeba dispar (paper I).

To determine the distribution of Blastocystis subtypes among Blastocystis- carriers in Sweden (paper II).

To describe the occurrence of intestinal parasites and genetic subtypes of Blastocystis sp. and Giardia intestinalis in diarrhoeic and non-diarrhoeic patients in Zanzibar, Tanzania (paper III).

To investigate the carriage of Blastocystis and Blastocystis subtypes and their relation to the faecal microbiota in Swedish university students before and after intercontinental travel (paper IV).

24 3. Methodological considerations

3.1 Paper I

In paper I, we performed a methodological evaluation of a complete diagnostic chain, including sample collection, sample storage, DNA extraction, and qPCR, to detect Entamoeba histolytica and Entamoeba dispar in faecal samples. Since the parts of this chain are dependent on each other we performed the evaluation in a series of experiments. For instance, a decreased positivity rate in the qPCR could be caused by a sampling error, DNA degeneration after sampling, low efficiency of the DNA extraction method, PCR inhibition, or inherent liabilities in the assay itself. The evaluated qPCR was a previously described duplex assay [51] with a common primer pair for both E. histolytica and E. dispar, and species specific Taqman probes for species determination (Table 2). This assay was chosen based on an evaluation by Qvarnstrom et al [245], in which it showed superior species specificity in comparison to two other qPCR assays. The analytic performance of the duplex qPCR was evaluated on DNA extracted from cultured trophozoites of E. histolytica strain HM-1:IMSS and of E. dispar SAW 760 as well as synthetic SSU-rDNA of both E. histolytica and E. dispar (Phthisis Diagnostics). An advantage of cultured trophozoites is that they are easily countable under the microscope, which can be utilised to create dilution series of known amounts of parasites to test detection limits. We also tested the detection of known amounts of trophozoites spiked in faecal samples. Synthetic SSU-rDNA is quantified by the manufacturer and is therefore even easier to use to for exact determination of the number of gene copies added to each reaction. The synthetic genes were used to evaluate the qPCR detection in samples containing varying amounts of SSU- rDNA from both Entamoeba species. Neither trophozoites nor synthetic DNA offers a real challenge for DNA extraction methods. Ideally, DNA extractions methods used in intestinal parasitology should be evaluated against the cyst form of parasites since the cellular components are sheltered by a rigid cyst wall. We therefore used four clinical samples containing cysts of E. dispar, confirmed by PCR, in the evaluation of DNA extraction methods. Three methods were evaluated: a manual guanidinium thiocyanate (Gua-SCN) based method, slightly modified from Boom et al [16], for its simplicity; the manual QIAamp DNA stool mini kit (Qiagen) for its wide use in molecular parasitology; and the automated Arrow Stool DNA kit (NorDiag), for its potential to limit hands on time in clinical diagnostics. Initial testing showed weaker performance with the Arrow Stool DNA kit, but with the inclusion of sample pre-treatment in

25 the form of bead beating with a FastPrep-b24 (MP Biomedicals), the method yielded results in line with the others. A limitation of the study is that we did not perform further evaluation of the DNA extraction methods on a larger set of clinical samples, including samples containing E. histolytica. However, a large-scale evaluation of DNA extraction methods was not in the scope of the study.

Table 2. Primers and probes of molecular detection methods used in papers I-III.

qPCR assays used for detection of intestinal parasites Assay type Species Primers and probes Ref.

Duplex E. histolytica/dispar Ehd-239F (5´-ATTGTCGTGGCATCCTAACTCA-3´) [51] qPCR Ehd-88R (5´-GCGGACGGCTCATTATAACA-3´) Entamoeba histolytica Probe histolytica 96T (VIC 5´-TCATTGAATGAATTGGCCATTT-3´-MGBNFQ) Entamoeba dispar Probe dispar 96T (FAM 5´-TTACTTACATAAATTGGCCACTTTG-3´-MGBNFQ)

Multiplex Giardia intestinalis Giardia 80F (5´-GACGGCTCAGGACAACGGTT-3´) [246] qPCR Giardia 127R (5´-TTGCCAGCGGTGTCCG-3´) Probe Giardia T (FAM-5´-CCCGCGGCGGTCCCTGCTAG-3´-BHQ1)

Cryptosporidium spp. CrF (5´-CGCTTCTCTAGCCTTTCATGA-3´) [247] CrR (5´-CTTCACGTGTGTTTGCCAAT-3´) Probe Crypto (CY5-5´-CCAATCACAGAATCATCAGAATCGACTGGTATC-3´-BHQ2)

Dientamoeba fragilis Df-124F (5´-CAACGGATGTCTTGGCTCTTTA-3´) [248] Df-221R (5´-TGCATTCAAAGATCGAACTTATCAC-3´) Probe Df-172revT (VIC-5´-CAATTCTAGCCGCTTAT-3´-BHQ1)

qPCR Blastocystis sp. Blasto Fwd F5 (5´-GGTCCGGTGAACACTTTGGATTT-3´) [227] Blasto Rev F2 (5´-CCTACGGAAACCTTGTTACGACTTCA-3´) Blasto Probe (FAM-5´-TCGTGTAAATCTTACCATTTAGAGGA-3´-MGBNFQ)

PCR assays used for subtyping of intestinal parasites Assay type Species Primers and probes Ref.

Sequencing Blastocystis sp. RD5 (5´-ATCTGGTTGATCCTGCCAGT-3´) [106] BhRDr (5´-GAGCTTTTTAACTGCAACAACG-3´)

PCR Giardia intestinalis Assemblage A Fwd (5´-AAAGAGATAGTTCGCGATGTC-3´) [248] assemblage A Assemblage A Rev (5´-ATTAACAAACAGGGAGACGTATG-3´) Giardia intestinalis Assemblage B Fwd (5´-GAAGTCATCTCTGGGGCAAG-3´) assemblage B Assemblage B Rev (5´-GAAGTCTAGATAAACGTGTCGG-3´)

To investigate the occurrence of PCR inhibition we used a linearized plasmid with a 720 bp insert of Drosophila melanogaster ALKcDNA in the vector pcDNA3 (Invitrogen) as an internal positive control (IPC). The IPC was detected by a separate qPCR described in paper I. The amount of IPC added to each DNA template was 125 copies per reaction tube and the expected Ct value was 29.3.

26 Sample fixatives containing formalin, such as SAF, that are used for light microscopy has been shown to interfere with PCR analysis [249]. Degradation of DNA can occur during transport of unpreserved faeces, and using ethanol as a fixative [250] or freezing faecal samples [251] has been recommended. In an effort to avoid DNA degradation during transport and facilitate DNA extraction we evaluated Gua-SCN for storage and transport. The influence of storage of faecal samples on the ability to detect Entamoeba DNA by qPCR was examined in different experiments using tenfold dilutions of E. histolytica and E. dispar trophozoites spiked in faeces. In the longest running experiment, the ability to detect E. dispar in spiked faecal samples stored in the Gua-SCN buffer L6T was compared to samples stored in phosphate buffered saline (PBS) during 48 days.

3.2 Paper II

In paper II, we performed subtyping of 68 clinical O&P samples from the Stockholm area that were positive for Blastocystis by microscopy, to determine the genetic diversity of Blastocystis in Sweden. Unfortunately, notifications from clinicians on travel history and symptomatology were only available in a small number of cases. This lack of patient data is a common concern for clinical microbiology laboratories in the analysis of patient samples. Nevertheless, the O&P exam is generally requested for patients with gastrointestinal symptoms or eosinophilia, and it seems unlikely that samples from completely asymptomatic individuals were included in the study. The samples were collected by the Karolinska University Hospital, Solna, Sweden prior to the evaluations performed in paper I, and divided into two parts, one part stored frozen and one part stored in 70% ethanol at +4 °C to limit DNA degradation. DNA was extracted from approximately 200 mg faeces using the QiaAMP DNA Stool Mini Kit (Qiagen). As per the manufacturer’s instructions, frozen samples were not thawed before extraction, but it was cumbersome to manually retrieve the frozen faeces from the sample tubes. Furthermore, several steps in the method required quite a lot of hands on time, making large scale DNA extractions with this method seem implausible. At the time of the study, there were two major methods used for subtyping of Blastocystis, the barcoding method described by Scicluna et al [106] and the sequence tagged site (STS) primers described by Yoshikawa et al [147]. We decided to use the barcoding method, in which subtype designation is achieved by sequencing of a ~600 bp segment of the SSU-rDNA. For Blastocystis subtyping, this is as specific as sequencing of the whole 1800 bp SSU-rDNA. The sequences obtained by the barcoding method were

27 compared by BLAST searches to previously deposited sequences in GenBank for subtype designation. This method offers the possibility to detect novel subtypes and to find discrete sequence differences within subtypes. Mixed carriage of more than one subtype represents a problem for the barcoding method because of preferential amplification of the most dominant subtype. The occurrence of double peaks in the chromatogram is a possible indication of mixed carriage, but the subtypes involved cannot be determined. The seven STS primers [147], that we chose not to use, were derived from RAPD analysis, and are each specific for STs 1-7. They offer the possibility to detect and resolve mixed carriage of subtypes. However, intra-subtype sequence variation, novel subtypes, and subtypes other than ST1-7 will go undetected. A low sensitivity of the STS primers to detect the most common variant of ST4 has also been reported [252]. The sequences we obtained with the barcoding method were assigned to subtypes and aligned with each other in search for sequence variations. Phylogenetic trees were constructed with the neighbor-joining method [253] to infer phylogenetic relationships among the sequences. Statistical analysis with the χ2 test was used for comparisons of subtype prevalence in different groups.

3.3 Paper III

In paper III, we investigated the presence of intestinal parasites, including Blastocystis subtypes, in people living in Jambiani village, Zanzibar, Tanzania. The sanitation conditions are generally poor in the area and local wells are used for water supply. Faecal samples were collected from 174 individuals that sought care at the health clinic in Jambiani. Among children 0-10 years 55/108 (51%) samples were from patients with diarrhoea and 53/108 (49%) from patients without diarrhoea. From patients over ten years of age samples were mainly obtained from those with diarrhoea, 62/66 (94%). Samples were collected in both SAF and the Gua-SCN buffer L2 (Paper I) for parasite detection with microscopy and molecular methods. Samples collected in SAF were concentrated by FECT prior to microscopy of both iodine stained wet smears and acid-fast stained smears, allowing for detection of cysts, ova, and oocysts of coccidian parasites. No routine for permanent staining of non-concentrated faecal samples was available at the laboratory so Dientamoeba fragilis or the trophozoite stage of other parasites were not detected by microscopy. DNA was extracted from faecal samples stored in L2-buffer by bead beating and automated extraction with the Arrow Stool DNA kit (paper I). PCR inhibition control was performed on all samples with the detection of 125 copies of the IPC as described in paper I. The duplex qPCR assay

28 evaluated in paper I was used to detect Entamoeba histolytica and Entamoeba dispar (Table 1). Giardia intestinalis, Dientamoeba fragilis, and Cryptosporidium spp. were detected by a multiplex qPCR, modified from an assay developed by the Dutch National Institute for Public Health and the Environment, using primers and probes previously described [245- 247] (Table 1). All samples were subjected to the Blastocystis barcoding PCR [106] (Table 1), and positive results were sequenced for subtype designation. Samples that were negative by the barcoding PCR were analysed with a Blastocystis qPCR developed by Stensvold et al [227] (Table 1) in an effort to increase the detection sensitivity for Blastocystis. The combination of these assays provides excellent coverage of medically important unicellular intestinal parasites. A possible addition, depending on the geographical setting, could be a PCR assay to detect Cyclospora cayetanensis [221]. A limitation of the study is that we did not include any detection methods for bacterial or viral enteropathogens. We observed a high detection rate of Giardia intestinalis by qPCR, and a method for Giardia assemblage typing was added post hoc, not only to determine genetic assemblages but also to validate the specificity of the qPCR. We chose a recently published conventional PCR, assay 4E1-HP, developed by Vanni et al [248], with specific primers for assemblage A and assemblage B (Table 1). This method was chosen because of is simplicity and the described excellent concurrence to results achieved by the more elaborate multi-locus genotyping of ß-giardin, TPI and GDH genes [248, 254]. The assay was originally presented as a duplex assay but we used the primer pairs in two separate reactions, which in our hands was more sensitive for mixed infections. Statistical analysis with the χ2 test was used for group comparisons of proportions of samples that were positive or negative by microscopy or PCR assays.

3.4 Paper IV

In paper IV, we performed metagenomic sequencing of faecal samples from 35 university students travelling abroad to determine the relationship between Blastocystis and the intestinal microbiota. Participants were included from a larger dataset of 99 students travelling for educational programs that were sampled before and after travel to determine the impact of travel on the occurrence of intestinal parasites, including Blastocystis subtypes (manuscript in preparation), and the acquisition of extended spectrum beta-lactamase (ESBL) producing bacteria [255]. Included in the metagenomic study were 35 health care students, 26/35 female, age 23-34 years, who travelled to Central Africa (17/35) or the Indian peninsula (18/35)

29 with a median travel duration of 34 days (range 14 to 150). The study design for paper IV offered an excellent opportunity to investigate to what extent Blastocystis is lost or acquired during travel, if the subtype distribution changes, and if changes in Blastocystis carriage is accompanied by changes in the bacterial microbiota. A temporal aspect could also be included in the analysis since the median travel duration was over one month. Collected faecal samples were frozen on arrival to the laboratory. The QIAamp DNA stool mini kit (Qiagen) was used for DNA extraction in samples included in the metagenomic analysis. Metagenomic sequencing was performed on a HiSeq 2000 (Illumina). The sequence data, consisting of 40 to 178 million filtered read pairs per sample, was previously used to study the impact of travel on intestinal carriage of bacterial antibiotic resistance genes [256] and had been deposited in the European Nucleotide Archive as project PRJEB7369. The read pairs were searched for SSU-rDNA sequences of both eukaryotes and prokaryotes using Metaxa2 2.0 [257]. The genus rDNA counts for each library were normalized to the total number of rDNA counts in that library, yielding relative abundances for each genus. SSU-rDNA sequences classified as deriving from the Blastocystis genus by Metaxa2 were compared to partial Blastocystis SSU-rDNA sequences available at the Blastocystis Sequence Typing database (http://pubmlst.org/blastocystis/) in October 2014. At the time, the database included 133 sequence variants belonging to STs 1-10. The presence of other subtypes would go undetected, but at the time of the study only STs 1-9 had been found in humans. To date, 3 human cases of ST12 from Bolivia [125] are the only observations of human carriage of subtypes other than STs 1-9. Furthermore, STs 1-4 represent over 94% of all human subtype observations. For these reasons, the limitation caused by the database selection is rather minor. The effect of Blastocystis and/or specific subtype presence or absence on total bacterial community composition was assessed on the family level using Metaxa2 Diversity Tools [258]. Spearman correlation was used to identify relationships between the relative abundances of individual bacterial genera and the total Blastocystis abundance or the abundance of the respective subtypes. P-values for correlations were adjusted for multiple testing using the Benjamini-Hochberg false discovery rate (FDR) [259] and tests with an FDR<0.05 were considered statistically significant. These correlations were calculated both before and after travel, as well as on all samples together and only correlations that were significant in all three cases were reported, as those seem to be stable both between individuals and over time. The mean bacterial genus richness and the relative abundance of specific bacterial genera were compared in Blastocystis positive and negative samples by using independent sample t-tests.

30 4. Results and discussion

4.1 The sensitivity of molecular detection methods

The duplex qPCR for E. histolytica and E. dispar evaluated in paper 1 showed very high detection sensitivity in several experiments, detecting for instance 5 genomes of E. histolytica per qPCR reaction, 1 trophozoite of E. histolytica per ml faecal suspension, and 5 synthetic copies of SSU-rDNA of either Entamoeba species per reaction. Using varying amounts of synthetic SSU-rDNA of both Entamoeba species in the same reaction we found that the efficiency to detect low amounts of DNA, 5-50 copies of SSU-rDNA, was impaired by the presence of the other amoeba in concentrations ≥ 500 gene copies per reaction. Thus, the limit of detection (LOD) for the non- dominating species in a mixed infection was between 50 and 500 gene copies per qPCR reaction. Since the SSU-rDNA in Entamoeba is located on an extra-chromosomal circular plasmid and present in about 200 copies per cell [260] this LOD corresponds to 0.25- 2.5 cells per reaction. This implies that the assays sensitivity is sufficient for resolving mixed infections in faecal samples from clinical cases. The detection sensitivity of PCR can also be evaluated in clinical samples by comparing its performance with that of microscopy. Such comparisons were done in the study of intestinal parasites in Zanzibar (paper III), bearing in mind that the sensitivity of microscopy would probably increase somewhat if three samples had been examined instead of only one. Detection rates for Blastocystis were 61% by PCR + qPCR and 20% by microscopy. Similarly, detection rates for Giardia intestinalis were 53% by qPCR and 14% by microscopy. Thus, PCR was more than three times as sensitive as microscopy for these two species. Three cases of Entamoeba histolytica/dispar were detected by microscopy, and these were identified as E. dispar by qPCR. Seven additional cases of E. dispar and one case of E. histolytica were found by qPCR in microscopy negative samples that would have been missed if only microscopy had been performed. Cryptosporidium spp. were only detected by qPCR in two individuals, both younger than two years. This unexpectedly low occurrence of Cryptosporidium could potentially be caused by problems in the DNA extraction or the qPCR assay. However, microscopy was only positive in one of these cases and no further cases of Cryptosporidium were detected by microscopy, indicating that the prevalence of this parasite was indeed low in our study population. A common concern from clinicians is that qPCR might be too sensitive for use in clinical diagnostics, possibly detecting minute number of parasites or detecting DNA from dead parasites. Detection of low amounts of DNA target,

31 as indicated by high Ct values, can present a problem in the management of patients, and further studies are needed to determine the clinical significance of such low positive samples. The possibility of detecting dead parasites may not be a major concern for non-invasive intestinal parasites. For G. intestinalis, a study from the Netherlands report clearance of Giardia DNA in all 75 included patients one week after successful treatment of giardiasis, confirming that qPCR can be used for treatment follow up [261]. In epidemiological studies of unicellular intestinal parasites, it is important to use molecular detections methods to accurately assess carrier rates. Molecular detection should also be used for follow-up in longitudinal studies and antimicrobial treatment trials, since the use of microscopy may lead to overestimations of parasite clearance rates. Molecular methods are also well suited for parasite detection in environmental samples, where the number of parasites per sample can be low.

4.2 PCR inhibition

PCR inhibition, resulting in false negative results or decreased amplification efficiency, was relatively common in the experiments in paper I that involved faecal material. Inhibition was clearly not an on-off phenomenon in our experiments and the use of an inhibitor control with known number of gene copies and expected Ct value in the qPCR was crucial to detect low-grade inhibition that resulted in false negative results in low positive samples. The straightforward approach of diluting the extracted DNA resolved the problem with inhibition. Since dilution also lowers the concentration of target DNA other measures to decrease the risk of inhibition should be explored.

4.3 Guanidinium thiocyanate as a transport medium

Gua-SCN is a chaotropic agent that causes cell lysis and inactivation of nucleases. It can also prevent DNA from being trapped in clinical sample residues, reducing the activity of PCR inhibitors [262]. Buffer solutions containing the chaotropic agent Gua-SCN was found to be suitable for sample storage for molecular diagnostics in our experiments in paper I. Trophozoites of E. dispar spiked in faeces and stored in a Gua-SCN buffer could be detected by qPCR with lower Ct values than trophozoites spiked in faeces that were stored in PBS. The buffer L2, with the lowest concentration of Gua-SCN, did not crystallize when refrigerated and this buffer was used in the collection of faecal samples in Zanzibar, Tanzania in paper III. In these 174 samples, total inhibition of the IPC (at 125 gene copies per reaction) was

32 seen in 11% (20/174). Dilution of the extracted DNA 1/10 resolved the inhibition in all cases. This can be compared to the samples collected before and after travel in the larger dataset of 99 university students from which participants were included in the study for paper IV. These 198 faecal samples were collected without additives, frozen on arrival to the laboratory, and subjected to the same DNA extraction protocol as used in paper III. Total inhibition of the IPC was seen in a staggering 68% (135/198) of the samples and total inhibition was still present in 5 samples after dilution of the DNA 1/10 (data not published). This indicates that using the Gua-SCN buffer L2 as a transport medium limits the occurrence of PCR inhibition in faecal samples that are sent for molecular parasite detection.

4.4 Blastocystis prevalence in Sweden and Zanzibar

In 2000, a Swedish study reported the prevalence of Blastocystis to be 4% in patients with diarrhoea and 9% in healthy controls [263]. However, this survey used microscopy for detection of Blastocystis and it is likely that this represents an underestimation of the true prevalence. There had been no studies using sensitive molecular methods to determine the prevalence of Blastocystis in Sweden prior to the work presented in this thesis. In paper IV, we found that 16/35 Swedish university students (46%) were positive for Blastocystis before intercontinental travel and 15/35 (43%) were positive after travel, based on the detection of Blastocystis SSU-rDNA in metagenomic data. This rather high occurrence of Blastocystis is in line with recent studies from Ireland [264], the Netherlands [265] and Denmark [218] that report Blastocystis carrier rates in healthy individuals of 56%, 33%, and 22% respectively, indicating that Blastocystis is common in the general population in Western Europe. In Zanzibar, Blastocystis sp. was the most commonly found intestinal parasite, detected in 106/174 (61%) study participants (paper III). Even higher detection rates, also using molecular detection methods, have been reported from Senegal (100%) [162], and recently also from Nigeria (84%) [161]. This confirms that Blastocystis is more common in low-income regions with low sanitation standards even as detection methods in epidemiological studies shifts from microscopy to PCR. We found Blastocystis to be equally prevalent in cases with and without diarrhoea (63% vs. 58%). As mentioned previously, this is a common finding in case- control studies of Blastocystis [209-212].

33 4.5 Blastocystis subtypes in Sweden and Zanzibar

The studies included in this thesis offer the first, and to date, only, descriptions of Blastocystis subtype occurrence in Sweden. Subtyping using the barcoding method [106] was successful in 63/68 clinical samples that were positive for Blastocystis by microscopy in paper II, and the subtype distribution was: ST1 16% (10/63), ST2 14% (9/63), ST3 48% (30/63), ST4 21% (13/63), and ST7 2% (1/63). Metagenomic sequencing was used to determine Blastocystis subtype carriage in 35 Swedish university students before and after travel in paper IV. Subtyping was successful for 14/16 Blastocystis carriers before travel, and the distribution of subtypes was: ST1 7% (1/14), ST2 14% (2/14), ST3 36% (5/14), ST4 43% (6/14). Combining the data from these two studies results in the following Swedish Blastocystis subtype distribution: ST1 14% (11/77), ST2 14% (11/77), ST3 45% (35/77), ST4 25% (19/77), and ST7 1% (1/77), which is similar to the subtype distribution found in Europe as shown in Fig. 4 (page 16). In the study of patients in Zanzibar (paper III), the initial barcoding PCR for Blastocystis sp. was positive in 85/174 (49%) individuals, and all these cases were successfully subtyped. An additional 21 cases of Blastocystis were detected by qPCR, and optimised attempts at subtyping these with the barcoding method were successful in seven out of 21. In total, the identified subtypes were: ST1 39% (36/92), ST2 30% (28/92), ST3 29% (27/92), and ST7 1% (1/92). The absence of ST4 is in line with the compiled data from Africa, in which ST4 represents 1% of the reported subtype observations (page 17, Fig. 5). We found no significant differences in carrier-rates in patients with and without diarrhoea for either ST1 (20% vs. 23%), ST2 (17% vs. 14%), or ST3 (15% vs. 16%). In paper II, we observed one possible case of mixed carriage with two or more subtypes, manifested as double peaks in the sequence chromatogram. No such observation was done in the study from Zanzibar (paper III). This does not entirely exclude the possibility of more cases with mixed subtype carriage in these two studies since there is a risk of preferential amplification of the dominant subtype when sequencing is performed directly from faeces. This was recently shown in a study by Scanlan et al [150] in which 50 Blastocystis samples that had previously been subtyped by sequencing were re-examined using subtype specific PCR, revealing previously undetected mixed subtype carriage in 22% of the samples. However, mixed subtype carriage was not seen among Swedish students in paper IV, although metagenomic sequencing is well suited to detect the sequences of more than one subtype. Further studies are needed to elucidate the true occurrence of mixed subtype carriage.

34 4.6 Giardia intestinalis in Zanzibar

Giardia intestinalis, a parasite included in WHO’s ‘Neglected Diseases Initiative’ [53], was highly prevalent in Zanzibar, detected by qPCR in 94/174 (54%) study participants (paper III). The presence of Giardia was high in all age groups investigated but was most prevalent in children two to five years of age, where the detection rate was 74%. No significant difference in prevalence was found between patients with and without diarrhoea (50% vs. 61%). The high prevalence of Giardia likely represents a significant health concern in Zanzibar, especially among children. In highly endemic settings, Giardia intestinalis is not only an important cause of acute diarrhoea but also the cause of a more chronic intestinal disease associated with malabsorption, malnutrition [266], stunted growth [267, 268], and even cognitive impairment [269, 270]. Even though treatment with antimicrobials such as metronidazole or tinidazole is effective, reinfections are common and other efforts are needed to reduce the disease burden. Such measures include access to clean water, good animal husbandry and adequate routines for disposal of manure and faeces [53]. The high detection rate of Giardia intestinalis by qPCR was confirmed by the conventional PCR assays for G. intestinalis assemblage A and B that gave positive results for 89 of 94 qPCR positive cases. The identified genetic assemblages were: assemblage A 7% (6/89), Assemblage B 89% (79/89), and assemblage A + B 4% (4/89). A predominance of assemblage B has also been observed on Pemba Island, Zanzibar, Tanzania [271] and Rwanda [272], as well as several other African countries, as described in paper III. Studies regarding the association of assemblage A and B to symptomatic infections have reached discordant conclusions [63-66]. Our data included too few cases of assemblage A infections to reliably investigate any differences in pathogenic potential between the assemblages. There is a need for even larger studies to determine if the non-dominant assemblage in Giardia endemic regions has a different virulence to the dominant assemblage.

4.7 Dientamoeba fragilis in Zanzibar

Dientamoeba fragilis was quite common in Zanzibar, detected by qPCR in 17% of the participants (paper III). We found no significant difference in prevalence of D. fragilis between patients with and without diarrhoea (15% vs. 21%).

35 4.8 The debated pathogenicity of Blastocystis

In paper III, we found no significant differences in carrier-rates in cases with and without diarrhoea for Blastocystis sp. or for the common Blastocystis STs 1-3. A lack of association between parasite carriage and diarrhoea was also seen for Giardia intestinalis and Dientamoeba fragilis. This is interesting in the context of Blastocystis debated pathogenicity since asymptomatic carriage of Giardia is common in areas of high endemicity, but Giardia is also an important cause of diarrhoea. In populations were enteropathogens are common, the results of case-control studies should not be used to rule out pathogenicity in the detected organisms. This was demonstrated in a case-control study on diarrhoea among children under five years in Tanzania, in which 52.2% of asymptomatic individuals carried one or more enteropathogens. The authors found that Shigella was the only detected organism statistically linked to diarrhoea – Salmonella, ETEC, EPEC, EAEC and Giardia were not [273]. Since Blastocystis is highly prevalent in most study settings, a high occurrence of asymptomatic carriage is not surprising. This does not exclude the possibility that Blastocystis can cause disease under certain conditions. Continued research in this field is important so that possible mechanisms behind symptomatic carriage of Blastocystis can be unravelled.

4.9 Age and gender distributions of Blastocystis

In contrast to what was found for Giardia intestinalis, the prevalence of Blastocystis in Zanzibar increased with increasing age, at least up to young adulthood (paper III). A Blastocystis prevalence increasing by age was also found in a study of children 2-14 years old in Nigeria [161]. Furthermore, a study from Libya reported that Blastocystis was more common in participants over 18 years than in those under 18 years [160]. An increasing prevalence with age might indicate a gradual establishment of Blastocystis carriage over time and that long-term colonization is common. In Zanzibar, we found no statistically significant difference in carrier-rates of Blastocystis between males (66%) and females (55%) (paper III). An observed difference in prevalence for ST2 in males (21.3%) compared to females (10%) was not statistical significant. Among Swedish Blastocystis cases analysed in paper II there was a marginally significant overrepresentation of ST3 in males (p = 0.049). An association between ST3 and males was also seen in a study from Qatar [177], in which the authors also found an association between ST1 and females. An association between ST1 and females was also seen in a study from Libya [160]. The sample sizes at the subtype level were quite small in these studies, and it remains to be

36 seen if these gender differences in carriage of ST1 and ST3 can be substantiated in future studies.

4.10 Temporal stability of Blastocystis carriage

In paper IV, we reported a high occurrence of Blastocystis in university students both before and after travel to Central Africa or the Indian peninsula, and no apparent impact of travel on the subtype distribution. Of the 16 individuals that were positive for Blastocystis before travel 13 were still positive after travel. Reliable subtype designation was achieved both before and after travel in ten of these and each of them carried the same Blastocystis subtype before and after travel. This suggests a persistence of a specific subtype in these individuals for the duration of their travel, which varied between three weeks and five months. This apparent stability in Blastocystis carriage was seen despite the quite drastic change in environment experienced by the travellers. To what extent Blastocystis causes long term colonization is still not fully investigated. In a study by Scanlan et al [264] four individuals consistently tested positive for the same Blastocystis strain during a study period of 6-10 years, while six individuals repeatedly tested negative for Blastocystis, confirming that colonization for several years can occur. However, larger longitudinal studies, using molecular detection methods, are needed to determine if Blastocystis predominately causes long lasting colonisation or shorter self-limiting carriage. Such studies could also be used to determine to what extent Blastocystis causes symptomatic episodes.

4.11 Blastocystis and the bacterial microbiota

In the metagenomic study of the intestinal microbiota presented in paper IV, we found no systematic significant differences in the bacterial compositions at the family level between Blastocystis positive and negative individuals or between carriers of different subtypes. We did, however, find associations between Blastocystis and certain bacterial genera. The relative abundance of Blastocystis was significantly positively correlated to the relative abundance of bacteria in the genera Sporolactobacillus (correlation coefficient 0.8731, p < 0.0001) and Candidatus Carsonella (correlation coefficient 0.6430, p < 0.0001), both before and after travel. When comparing Blastocystis positive samples with negative samples we found positive samples to be associated with an increased relative abundance of Sporolactobacillus (p = 0.00021). An increased relative abundance of Candidatus Carsonella (p = 0.038) was also

37 noted for Blastocystis positive samples in the total material, however, analysis of samples taken before and after travel separately did not reach significance (p = 0.19, and p = 0.097, respectively). Sporolactobacillus are anaerobic, spore-forming, and lactic acid- producing bacteria [274, 275], that have been mostly isolated from environmental sources such as soil and decaying foodstuffs [276-278]. Not much is known about its role in the intestinal microbiota, but the type species, Sporolactobacillus inulinus, has been suggested as a possible probiotic species [279]. In general, intestinal lactic acid bacteria are favoured by diets high in non-metabolizable sugars which are typically found in vegetables [280]. The association between Blastocystis and Sporolactobacillus might suggest that they have similar transmission sources, that they share the same ecological requirements in the intestine and/or that they are favoured by similar diets. The association between Blastocystis and Candidatus Carsonella is a curious finding. Candidatus Carsonella ruddii, the only described species of this genus, is a strict endosymbiont of sap feeding insects [281], and has a tiny genome that seems to lack genes essential for bacterial life on its own [282]. Bacteria-like endosymbionts of Blastocystis have been described [283], and Blastocystis could perhaps be associated with some other, yet undescribed, species of the Candidatus Carsonella genus. Another possibility is that the bacteria have been ingested through food contaminated with insect cells. The relative abundance of Bacteroides was significantly lower in faecal samples containing Blastocystis (p = 0.00027). A similar negative association between Blastocystis and Bacteroides has been found in two studies from Denmark, one using metagenomics [143] and one using qPCR quantification of major bacterial taxa [284]. The Bacteroides-driven enterotype has previously been linked to diets high in protein and animal fat, typical of Western diet [285, 286], and it is possible that Blastocystis carriage is favoured by a different diet. Both the positive correlation to Sporolactobacillus and the negative correlation to Bacteroides could indirectly imply a link between Blastocystis carriage and intake of vegetables. Blastocystis-carriers also had a significantly higher richness of bacterial genera compared to non-carriers (p = 0.0022). This strengthens the association between Blastocystis and high bacterial diversity that has been reported in two other metagenomic studies [143, 287]. Since a gut microbiota with high diversity is generally considered as a healthy microbiota, Blastocystis might be seen as an indicator of gastrointestinal health, an opinion also voiced by other researchers [143, 264, 287]. However, the gastrointestinal microbiota is a complex and dynamic ecological community in which the microorganisms interact both with each

38 other and the human host. A clear definition of the healthy human gut microbiota is still lacking. Diversity and stability are regarded as important features but the early hypothesis of a healthy core composition of prevalent microorganisms has been debated and been replaced by the hypothesis that special biomolecular functions produced by the microorganisms is what constitutes a healthy microbiota [288-290]. Further studies are needed to elucidate if Blastocystis has a role in maintaining or deteriorating a healthy balance in the intestinal ecosystem.

4.12 Blastocystis ST4

In the metagenomic study presented in paper IV, we compared the two most commonly found subtypes, ST3 and ST4, and found that the negative association between Blastocystis and Bacteroides was significant for ST4 (p = 0.0057) but not for ST3 (p = 0.73). The association between Blastocystis and high richness of bacterial genera was also largely driven by ST4 (p = 0.000037) and not by ST3 (p = 0.71). Although the sample size at the subtype level was small, and inter-individual differences could have influenced the results, the prospect of ST4 having specific associations to the intestinal microbiota is very interesting, especially considering that ST4 also has other unique traits. In paper II, we found intra-subtype sequence variation between strains in the ~600 bp barcoding region for STs 1-3, but no intra-subtype sequence variation in the 13 cases of ST4. We also found that the SSU-rDNA sequences of ST4 previously deposited in Genbank separated into only two phylogenetic clades, Clade 1 and 2. In an MLST study of genes coding for the mitochondrion-like organelle in Blastocystis ST3 and ST4, Stensvold et al. [291] concluded that 98% of the human ST4 sequences available in GenBank belonged to Clade 1. The MLST results from that study confirmed the limited genetic diversity in ST4 as only 5 sequence types were identified in 50 ST4s belonging to clade 1 compared to the 58 sequence types that were identified in 81 isolates of ST3. As we put forth in paper II, this limited genetic diversity within ST4 could indicate that the ST4 lineage has expanded more recently on an evolutionary scale. This is interesting in the context of the limited geographical distribution of ST4. To date, ST4 is the second most common subtype in Europe and the third most common subtype in Australia, but rare in other parts of the world. The cause of this is unknown but could possibly be explained by differences in host factors or differences in exposure to ST4, perhaps through diet, water supply or animal contact. There is a vast geographical distance between Europe and Australia but both are industrialised regions with high levels of sanitation. A typical western diet is

39 also common in both regions. It is therefore somewhat surprising that ST4 was negatively associated to Bacteroides, since enrichment in Bacteroides has been linked to western diets. Perhaps diet is not the decisive factor and carriage of ST4 is instead reliant on genetic host factors. The limited genetic diversity in ST4 could speculatively indicate an evolutionary cospeciation between ST4 and people of European descent that was developed sometime before Europeans colonised Australia. If this is true, ST4 would also be common in the USA since the majority of the US population have European ancestry [292]. Unfortunately, there is only sparse data on Blastocystis subtypes in the USA, and the occurrence of ST4 remains to be determined. The hypothesis that ST4 carriage is favoured by genetic host factors could also be tested by comparing the occurrence of ST4 in different ethnic groups in industrialised regions. So far, the reasons behind the limited geographic spread of ST4 is not known and represents and interesting topic for future studies.

40 5. Conclusions

PCR-based methods were highly sensitive for detection of unicellular intestinal parasites, but could be partially or completely inhibited by substances present in faeces. A sample transport medium containing guanidinium thiocyanate was shown to limit the occurrence of PCR inhibition.

The prevalence of Blastocystis in Swedish university students was over 40%, which is markedly higher than what was previously estimated. Blastocystis ST3 and ST4 were the two most commonly found Blastocystis subtypes in Sweden, which is similar to results from other European countries.

Blastocystis sp. and Giardia intestinalis were both commonly detected in Zanzibar, each with a prevalence exceeding 50%. While G. intestinalis was most common in the ages 2-5 years, the prevalence of Blastocystis increased with increasing age, at least up to young adulthood. Neither Blastocystis nor G. intestinalis could be statistically linked to diarrhoea by case-control analysis. Blastocystis ST1, ST2, and ST3 were common in Zanzibar, but no carriage of ST4 was found.

A persistence of Blastocystis subtypes was seen in Swedes that travelled to low-resource regions, indicating stability in Blastocystis carriage.

Metagenomic sequencing of faecal samples from Swedes revealed that Blastocystis was positively associated with intestinal bacterial genus richness, possibly signifying gastrointestinal health. Blastocystis was also positively associated with the bacterial genera Sporolactobacillus and Candidatus Carsonella, and negatively associated with the genus Bacteroides.

Blastocystis ST4 was shown to have limited intra-subtype genetic diversity and limited geographic spread. ST4 was also found to be the major driver behind the positive association between Blastocystis and bacterial genus richness and the negative association with Bacteroides.

41 6. Acknowledgements

I would like to thank everyone that has contributed to this thesis, both study participants and fellow researchers. I would especially like to acknowledge the following persons:

Birgitta Evengård, my supervisor, thank you for introducing me to the field of parasitology, creating the opportunity for me to study intestinal parasites, and guiding and supporting me throughout the process of my doctoral studies.

Margareta Granlund, my co-supervisor, thank you for all the help and encouragement you have provided over these years. I am impressed not only by your knowledge in parasitology and bacteriology but also that you always seem to have time for questions and discussions.

Graham Clark, thank you for taking me on in your laboratory, introducing me to the world of Entamoeba and Blastocystis, and all the help you have provided during the years.

Christen Rune Stensvold, thank you for all the support you have provided in Blastocystis subtyping. I have followed your research blog on Blastocystis with keen interest.

Johan Bengtsson-Palme, thank you for fruitful discussions and excellent collaboration in the metagenomics project. Maybe we will get the chance to meet in real life someday.

Martin Angelin, thank you for great discussions and collaborations during our parallel doctoral studies.

Anders Johansson, thank you for initiating the metagenomic study and your constructive feedback all through my studies.

Helén Edebro, always cheerful and effective in laboratory, thank you for all the work you have put in during these years.

Satu Koskiniemi, thank you for helping me with the molecular side of things.

Ida Hedberg, thank you for the laboratory work you performed on Entamoeba.

42 Thank you, Linn Samuelsson and Sara Viksten, you were both excellent students that showed great understanding of molecular parasitology.

Thank you, Lillemor Karlsson, Christina Seeth Grönfors, Christine Stenström, and Anna Dyrsmeds, for the work you put in by collecting Blastocystis samples.

Omar Hamsa, thank you for managing the sample collection in Zanzibar.

Thank you, Titia Kortbeek and Jeroen Roelfsema, for sharing your multiplex PCR and a pleasant dinner in Utrecht.

Thank you, Maria Casserdahl, Marianne Lebbad, Agheleh Hosseiny, Martin Ferm, and Johan Holmlund, for invaluable contributions to the studies.

Thank you, Urban Kumlin and all my colleagues at the diagnostic laboratory, for letting me spend time with my research.

My good friend, Marcus Forsblad, thank you for fun times in Amiens and Monte Grappa.

Agneta and Per-Olof, thank you for helping out with the kids.

Mum, dad, and sis, thank you for supporting me.

My kids, Viggo and Sofie, I love you so much, thank you for all the love you give me.

Sara, my lovely wife. What would I be without you? Thank you for all the support and positivity you have given me through all the years we have been together. I look forward to the rest of our lives together. A relaxing vacation would be a nice start, don’t you think?

43 7. References

1. Cox FE. History of human parasitology. Clin Microbiol Rev. 2002;15:595-612.

2. Ford BJ. The Discovery of Giardia. Microscope 2005 53, 147-53

3. Lesh FA. Massive development of amebas in the large intestine. Fedor Aleksandrovich Lesh (Lösch). Am J Trop Med Hyg. 1975;24:383-92.

4. Yang J, Scholten T. A fixative for intestinal parasites permitting the use of concentration and permanent staining procedures. Am J Clin Pathol. 1977;67:300-4.

5. Mank TG, Zaat JO, Blotkamp J, Polderman AM. Comparison of fresh versus sodium acetate acetic acid formalin preserved stool specimens for diagnosis of intestinal protozoal infections. Eur J Clin Microbiol Infect Dis. 1995 c;14:1076-81.

6. Allen AVH, Ridley DS. Further observations on the formol-ether concentration technique for faecal parasites. J Clin Pathol. 1970;23:545-6.

7. Wheatley WB. A rapid staining procedure for intestinal amoebae and flagellates. Am J Clin Pathol. 1951;21:990-1.

8. Heidenhain M. Neue Untersuchungen über die Centralkörper und ihre Beziehungen zum Kern- und Zellenprotoplasma. Arch Mikr Anat. 1894;43:423–758.

9. Kohn J. A one stage permanent staining method for fecal protozoa. Dapim Refu. 1960;19:160.

10. van Gool T, Weijts R, Lommerse E, Mank TG. Triple Faeces Test: an effective tool for detection of intestinal parasites in routine clinical practice. Eur J Clin Microbiol Infect Dis. 2003;22:284-90.

11. Ziehl F. Zur Farbung des Tuberkelbacillus. Deutsche Med Wochenschr. 1882;8:451.

12. Neelsen F. Ein Casuistischer Beitrag zur Lehre von der Tuberkulose. Zentrablau für die medizinischen Wissenschafen. 1883;21:497-501.

13. Cartwright CP. Utility of multiple-stool specimen ova and parasite examinations in a high prevalence setting. J Clin Microbiol. 1999;6:2408-11.

14. Marti H, Koella JC. Multiple stool examinations for ova and parasites and rate of false- negative results. J Clin Microbiol. 1993;6:3044-5.

15. Chomczynski P, Sacchi N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem. 1987;162:156-9.

16. Boom R, Sol CJ, Salimans MM, Jansen CL, Wertheim-van Dillen PM, van der Noordaa J. Rapid and simple method for purification of nucleic acids. J Clin Microbiol. 1990;28:495- 503.

17. Tan SC, Yiap BC. DNA, RNA, and protein extraction: the past and the present. J Biomed Biotechnol. 2009;2009:574398.

18. Mullis K, Faloona F, Scharf S, Saiki R, Horn G, Erlich H. Specific enzymatic amplification of DNA in vitro: the polymerase chain reaction. 1986. Biotechnology. 1992;24:17-27.

19. Higuchi R, Dollinger G, Walsh PS, Griffith R. Simultaneous amplification and detection of specific DNA sequences. Biotechnology (N Y). 1992;10:413-7.

44 20. Holland PM, Abramson RD, Watson R, Gelfand DH. Detection of specific polymerase chain reaction product by utilizing the 5'----3' exonuclease activity of Thermus aquaticus DNA polymerase. Proc Natl Acad Sci U S A. 1991;88:7276-80.

21. Verweij JJ, Stensvold CR. Molecular testing for clinical diagnosis and epidemiological investigations of intestinal parasitic infections. Clin Microbiol Rev. 2014;27:371-418.

22. van Lieshout L, Roestenberg M2. Clinical consequences of new diagnostic tools for intestinal parasites. Clin Microbiol Infect. 2015;21:520-8.

23. Roberts RJ. Restriction endonucleases. CRC Crit Rev Biochem. 1976;4:123-64.

24. Clark CG. Riboprinting: a tool for the study of genetic diversity in microorganisms. J Eukaryot Microbiol. 1997;44:277-83.

25. Welsh J, McClelland M. Fingerprinting genomes using PCR with arbitrary primers. Nucleic Acids Res. 1990;18:7213-8.

26. Prober JM, Trainor GL, Dam RJ, Hobbs FW, Robertson CW, Zagursky RJ, et al. A system for rapid DNA sequencing with fluorescent chain-terminating dideoxynucleotides. Science. 1987;238:336-41.

27. Maiden MC, Bygraves JA, Feil E, Morelli G, Russell JE, Urwin R, et al. Multilocus sequence typing: a portable approach to the identification of clones within populations of pathogenic microorganisms. Proc Natl Acad Sci USA. 1998;95:3140-5

28. Jex AR, Pangasa A, Campbell BE, Whipp M, Hogg G, Sinclair M, et al. Classification of Cryptosporidium species from patients with sporadic cryptosporidiosis by use of sequence-based multilocus analysis following mutation scanning. J Clin Microbiol. 2008;46:2252-62.

29. Handelsman J, Rondon MR, Brady SF, Clardy J, Goodman RM. Molecular biological access to the chemistry of unknown soil microbes: a new frontier for natural products. Chem Biol. 1998;5:R245-9.

30. Cho I, Blaser MJ. The human microbiome: at the interface of health and disease. Nat Rev Genet. 2012;13:260-70.

31. Parfrey LW, Walters WA, Lauber CL, Clemente JC, Berg-Lyons D, Teiling C, et al. Communities of microbial eukaryotes in the mammalian gut within the context of environmental eukaryotic diversity. Front Microbiol. 2014;5:298

32. Bäckhed F, Fraser CM, Ringel Y, Sanders ME, Sartor RB, Sherman PM et al. Defining a healthy human gut microbiome: current concepts, future directions, and clinical applications. Cell Host Microbe. 2012;12:611-22.

33. de Vos WM, de Vos EA. Role of the intestinal microbiome in health and disease: from correlation to causation. Nutr Rev. 2012;70 Suppl 1:S45-56

34. Arumugam M, Raes J, Pelletier E, Le Paslier D, Yamada T, Mende DR et al. Enterotypes of the human gut microbiome. Nature. 2011;473:174-80.

35. Lozupone CA, Stombaugh J, Gonzalez A, Ackermann G, Wendel D, Vázquez-Baeza Y, et al. Meta-analyses of studies of the human microbiota. Genome Res. 2013;23:1704-14.

36. Brown CT, Davis-Richardson AG, Giongo A, Gano KA, Crabb DB, Mukherjee N, et al. Gut microbiome metagenomics analysis suggests a functional model for the development of autoimmunity for type 1 diabetes. PLoS One. 2011;6:e25792.

45 37. Larsen N, Vogensen FK, van den Berg FW, Nielsen DS, Andreasen AS, Pedersen BK, et al. Gut microbiota in human adults with type 2 diabetes differs from non-diabetic adults. PLoS One. 2010;5:e9085.

38. Karlsson FH, Fåk F, Nookaew I, Tremaroli V, Fagerberg B, Petranovic D et al. Symptomatic atherosclerosis is associated with an altered gut metagenome. Nat Commun. 2012;3:1245.

39. Turnbaugh PJ, Hamady M, Yatsunenko T, Cantarel BL, Duncan A, Ley RE, et al. A core gut microbiome in obese and lean twins. Nature. 2009;457:480-4.

40. Musso G, Gambino R, Cassader M. Interactions between gut microbiota and host metabolism predisposing to obesity and diabetes. Annu Rev Med. 2011;62:361-80.

41. Kaser A, Zeissig S, Blumberg RS. Inflammatory bowel disease. Annu Rev Immunol. 2010;28:573-621.

42. Lepage P, Häsler R, Spehlmann ME, Rehman A, Zvirbliene A, Begun A, et al. Twin study indicates loss of interaction between microbiota and mucosa of patients with ulcerative colitis. Gastroenterology. 2011;141:227-36.

43. Midtvedt T, Zabarovsky E, Norin E, Bark J, Gizatullin R, Kashuba V, et al. Increase of faecal tryptic activity relates to changes in the intestinal microbiome: analysis of Crohn's disease with a multidisciplinary platform. PLoS One. 2008;8:e66074.

44. Carroll IM, Ringel-Kulka T, Siddle JP, Ringel Y. Alterations in composition and diversity of the intestinal microbiota in patients with diarrhea-predominant irritable bowel syndrome. Neurogastroenterol Motil. 2012;24:521-30.

45. Upadhyaya B, McCormack L, Fardin-Kia AR, Juenemann R, Nichenametla S, Clapper J, et al. Impact of dietary resistant starch type 4 on human gut microbiota and immunometabolic functions. Sci Rep. 2016;6:28797.

46. Koh A, De Vadder F, Kovatcheva-Datchary P, Bäckhed F. From dietary fiber to host physiology: short-chain fatty acids as key bacterial metabolites. Cell. 2016;165:1332-45.

47. Adl SM, Simpson AG, Lane CE, Lukeš J, Bass D, Bowser SS, et al. The revised classification of eukaryotes. J Eukaryot Microbiol. 2012;59:429-93.

48. World Health Organisation. Amoebiasis. Wkly Epidemiol Rec. 1997;72:97-9

49. Stanley SL Jr. Amoebiasis. Lancet. 2003;361:1025-34.

50. Diamond LS, Clark CG. A redescription of Entamoeba histolytica Schaudinn, 1903 (Emended Walker, 1911) separating it from Entamoeba dispar Brumpt, 1925. J Eukaryot Microbiol. 1993;40:340-4.

51. Verweij JJ, Oostvogel F, Brienen EA, Nang-Beifubah A, Ziem J, Polderman AM. Short communication: Prevalence of Entamoeba histolytica and Entamoeba dispar in northern Ghana. Trop Med Int Health. 2003;8:1153-6.

52. Abramowicz M, editor. Drugs for Parasitic Infections. New Rochelle, NY: The Medical Letter, Inc. 2013;11 Suppl:e1-31.

53. Savioli L, Smith H, Thompson A. Giardia and Cryptosporidium join the 'Neglected Diseases Initiative'. Trends Parasitol. 2006;22:203-8.

54. Folkhälsomyndigheten. www.folkhalsomyndigheten.se/folkhalsorapportering-statistik/ Accessed 16 feb 2017.

46 55. Ljungström I, Castor B. Immune response to Giardia lamblia in a water-borne outbreak of giardiasis in Sweden. J Med Microbiol. 1992;36:347-52.

56. Nygård K, Schimmer B, Søbstad Ø, Walde A, Tveit I, Langeland N, et al. A large community outbreak of waterborne giardiasis-delayed detection in a non-endemic urban area. BMC Public Health. 2006;6:141.

57. Adam EA, Yoder JS, Gould LH, Hlavsa MC, Gargano JW. Giardiasis outbreaks in the United States, 1971-2011. Epidemiol Infect. 2016;144:2790-801.

58. Enserink R, van den Wijngaard C, Bruijning-Verhagen P, van Asten L, Mughini-Gras L, Duizer E, et al. Gastroenteritis attributable to 16 enteropathogens in children attending day care: significant effects of rotavirus, norovirus, astrovirus, Cryptosporidium and Giardia. Pediatr Infect Dis J. 2015;34:5-10.

59. Feng Y, Xiao L. Zoonotic potential and molecular epidemiology of Giardia species and giardiasis. Clin Microbiol Rev. 2011;24:110-40.

60. Heyworth MF. Giardia duodenalis genetic assemblages and hosts. Parasite. 2016;23:13.

61. Fantinatti M, Bello AR, Fernandes O, Da-Cruz AM. Identification of Giardia lamblia Assemblage E in Humans Points to a New Anthropozoonotic Cycle. J Infect Dis. 2016;214:1256-9.

62. Franzén O, Jerlström-Hultqvist J, Castro E, Sherwood E, Ankarklev J, Reiner DS, et al. Draft genome sequencing of Giardia intestinalis assemblage B isolate GS: is human giardiasis caused by two different species? PLoS Pathog. 2009;5:e1000560.

63. Haque R, Roy S, Kabir M, Stroup SE, Mondal D, Houpt ER. Giardia assemblage A infection and diarrhea in Bangladesh. J Infect Dis. 2005;192:2171-3.

64. Pestechian N, Rasekh H, Rostami-Nejad M, Yousofi HA, Hosseini-Safa A. Molecular identification of Giardia lamblia; is there any correlation between diarrhea and genotyping in Iranian population? Gastroenterol Hepatol Bed Bench. 2014;7:168-72.

65. ElBakri A, Samie A, Bessong P, Potgieter N, Odeh RA. Detection and molecular characterisation of Giardia lamblia genotypes in Sharjah, United Arab Emirates. Trans R Soc Trop Med Hyg. 2014;108:466-73.

66. Puebla LJ, Núñez FA, Fernández YA, Fraga J, Rivero LR, Millán IA, et al. Correlation of Giardia duodenalis assemblages with clinical and epidemiological data in Cuban children. Infect Genet Evol. 2014;23:7-12.

67. Šlapeta J. Cryptosporidiosis and Cryptosporidium species in animals and humans: a thirty colour rainbow? Int J Parasitol. 2013;43:957-70.

68. Kotloff KL, Nataro JP, Blackwelder WC, Nasrin D, Farag TH, Panchalingam S, et al. Burden and aetiology of diarrhoeal disease in infants and young children in developing countries (the Global Enteric Multicenter Study, GEMS): a prospective, case-control study. Lancet. 2013;382:209-22.

69. Centers for Disease Control and Prevention. www.cdc.gov/parasites/crypto/pathogen.html Accessed 16 feb 2017.

70. Mac Kenzie WR, Hoxie NJ, Proctor ME, Gradus MS, Blair KA, Peterson DE, et al. A massive outbreak in Milwaukee of Cryptosporidium infection transmitted through the public water supply. N Engl J Med. 1994;331:161-7.

71. Widerström M, Schönning C, Lilja M, Lebbad M, Ljung T, Allestam G, et al. Large outbreak of Cryptosporidium hominis infection transmitted through the public water supply, Sweden. Emerg Infect Dis. 2014;20:581-9.

47 72. Rehn M, Wallensten A, Widerström M, Lilja M, Grunewald M, Stenmark S, et al. Post- infection symptoms following two large waterborne outbreaks of Cryptosporidium hominis in Northern Sweden, 2010-2011. BMC Public Health. 2015;15:529.

73. Coupe S, Sarfati C, Hamane S, Derouin F. Detection of Cryptosporidium and identification to the species level by nested PCR and restriction fragment length polymorphism. J Clin Microbiol. 2005;43:1017-23.

74. Peng MM, Matos O, Gatei W, Das P, Stantic-Pavlinic M, Bern C, et al. A comparison of Cryptosporidium subgenotypes from several geographic regions. J Eukaryot Microbiol. 2001;Suppl:28S-31S.

75. Alves M, Xiao L, Sulaiman I, Lal AA, Matos O, Antunes F. Subgenotype analysis of Cryptosporidium isolates from humans, cattle, and zoo ruminants in Portugal. J Clin Microbiol. 2003;41:2744-7.

76. Rossignol JF, Ayoub A, Ayers MS. Treatment of diarrhea caused by Cryptosporidium parvum: a prospective randomized, double-blind, placebo-controlled study of Nitazoxanide. J Infect Dis. 2001;184:103-6.

77. Amadi B, Mwiya M, Sianongo S, Payne L, Watuka A, Katubulushi M, Kelly P. High dose prolonged treatment with nitazoxanide is not effective for cryptosporidiosis in HIV positive Zambian children: a randomised controlled trial. BMC Infect Dis. 2009 Dec 2;9:195.

78. Kean BH, Malloch CL. The neglected ameba: Dientamoeba fragilis. A report of 100 "pure" infections. Am J Dig Dis. 1966;11:735-46.

79. Spencer MJ, Garcia LS, Chapin MR. Dientamoeba fragilis. An intestinal pathogen in children? Am J Dis Child. 1979;133:390-3.

80. Norberg A, Nord CE, Evengård B. Dientamoeba fragilis--a protozoal infection which may cause severe bowel distress. Clin Microbiol Infect. 2003;9:65-8.

81. Ögren J, Dienus O, Löfgren S, Einemo IM, Iveroth P, Matussek A. Dientamoeba fragilis prevalence coincides with gastrointestinal symptoms in children less than 11 years old in Sweden. Eur J Clin Microbiol Infect Dis. 2015;34:1995-8.

82. Bruijnesteijn van Coppenraet LE, Dullaert-de Boer M, Ruijs GJ, van der Reijden WA, van der Zanden AG, Weel JF, Schuurs TA. Case-control comparison of bacterial and protozoan microorganisms associated with gastroenteritis: application of molecular detection. Clin Microbiol Infect. 2015;21:592.e9-19.

83. de Jong MJ, Korterink JJ, Benninga MA, Hilbink M, Widdershoven J, Deckers-Kocken JM. Dientamoeba fragilis and chronic abdominal pain in children: a case-control study. Arch Dis Child. 2014;99:1109-13.

84. Stark D, Barratt J, Chan D, Ellis JT. Dientamoeba fragilis, the Neglected Trichomonad of the Human Bowel. Clin Microbiol Rev. 2016;29:553-80.

85. Röser D, Nejsum P, Carlsgart AJ, Nielsen HV, Stensvold CR. DNA of Dientamoeba fragilis detected within surface-sterilized eggs of Enterobius vermicularis. Exp Parasitol. 2013;133:57-61.

86. Ögren J, Dienus O, Löfgren S, Iveroth P, Matussek A. Dientamoeba fragilis DNA detection in Enterobius vermicularis eggs. Pathog Dis. 2013;69:157-8.

87. Stark D, Garcia LS, Barratt JL, Phillips O, Roberts T, Marriott D, et al. Description of Dientamoeba fragilis cyst and precystic forms from human samples. J Clin Microbiol. 2014;52:2680-3.

48 88. Windsor JJ, Johnson EH. Dientamoeba fragilis: the unflagellated human flagellate. Br J Biomed Sci. 1999;56:293-306.

89. Windsor JJ, Clark CG, Macfarlane L. Molecular typing of Dientamoeba fragilis. Br J Biomed Sci. 2004;61:153-5.

90. Nagata N, Marriott D, Harkness J, Ellis JT, Stark D. Current treatment options for Dientamoeba fragilis infections. Int J Parasitol Drugs Drug Resist. 2012;2:204-15.

91. Alexieff A. Sur la neture des formations dites ”Kystes de Trichomonas intestinalis”. C R Soc Biol. 1911;71:296-8.

92. Stensvold CR, Nielsen HV, Mølbak K, Smith HV. Pursuing the clinical significance of Blastocystis – diagnostic limitations. Trends Parasitol 2009;25:23-29

93. Tan KS. New insights on classification, identification, and clinical relevance of Blastocystis spp. Clin Microbiol Rev. 2008;21:639-65.

94. Silberman JD, Sogin ML, Leipe DD, Clark CG. Human parasite finds taxonomic home. Nature. 1996;380:398.

95. Krajaejun T, Sathapatayavongs B, Pracharktam R, Nitiyanant P, Leelachaikul P, Wanachiwanawin W, et al. Clinical and epidemiological analyses of human pythiosis in Thailand. Clin Infect Dis. 2006;43:569-76.

96. Stensvold CR, Suresh GK, Tan KS, Thompson RC, Traub RJ, Viscogliosi E, et al. Terminology for Blastocystis subtypes--a consensus. Trends Parasitol. 2007;23:93-6.

97. Clark CG. Extensive genetic diversity in Blastocystis hominis. Mol Biochem Parasitol. 1997;87:79-83.

98. Böhm-Gloning B, Knobloch J, Walderich B. Five subgroups of Blastocystis hominis from symptomatic and asymptomatic patients revealed by restriction site analysis of PCR- amplified 16S-like rDNA. Trop Med Int Health. 1997;2:771-8.

99. Yoshikawa H, Nagano I, Wu Z, Yap EH, Singh M, Takahashi Y. Genomic polymorphism among Blastocystis hominis strains and development of subtype-specific diagnostic primers. Mol Cell Probes. 1998;12:153-9.

100. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000;38:1324-30.

101. Kaneda Y, Horiki N, Cheng XJ, Fujita Y, Maruyama M, Tachibana H. Ribodemes of Blastocystis hominis isolated in Japan. Am J Trop Med Hyg. 2001;65:393-6.

102. Yoshikawa H, Wu Z, Nagano I, Takahashi Y. Molecular comparative studies among Blastocystis isolates obtained from humans and animals. J Parasitol. 2003;89:585-94.

103. Arisue N, Hashimoto T, Yoshikawa H. Sequence heterogeneity of the small subunit ribosomal RNA genes among Blastocystis isolates. Parasitology. 2003;126:1-9.

104. Noël C, Peyronnet C, Gerbod D, Edgcomb VP, Delgado-Viscogliosi P, Sogin ML, et al. Phylogenetic analysis of Blastocystis isolates from different hosts based on the comparison of small-subunit rRNA gene sequences. Mol Biochem Parasitol. 2003;126:119-23.

105. Noël C, Dufernez F, Gerbod D, Edgcomb VP, Delgado-Viscogliosi P, Ho LC, et al. Molecular phylogenies of Blastocystis isolates from different hosts: implications for genetic diversity, identification of species, and zoonosis. J Clin Microbiol. 2005;43:348-55.

106. Scicluna SM, Tawari B, Clark CG. DNA barcoding of Blastocystis. Protist. 2006;157:77-85.

49 107. Stensvold CR, Alfellani MA, Nørskov-Lauritsen S, Prip K, Victory EL, Maddox C, et al. Subtype distribution of Blastocystis isolates from synanthropic and zoo animals and identification of a new subtype. Int J Parasitol. 2009;39:473-9.

108. Parkar U, Traub RJ, Vitali S, Elliot A, Levecke B, Robertson I, Geurden T, Steele J, Drake B, Thompson RC. Molecular characterization of Blastocystis isolates from zoo animals and their animal-keepers. Vet Parasitol. 2010;169:8-17.

109. Fayer R, Santin M, Macarisin D. Detection of concurrent infection of dairy cattle with Blastocystis, Cryptosporidium, Giardia, and Enterocytozoon by molecular and microscopic methods. Parasitol Res. 2012;111:1349-55.

110. Alfellani MA, Taner-Mulla D, Jacob AS, Imeede CA, Yoshikawa H, Stensvold CR, Clark CG. Genetic diversity of Blastocystis in livestock and zoo animals. Protist. 2013;164:497-509.

111. Stenzel DJ, Boreham PF. Blastocystis hominis revisited. Clin Microbiol Rev. 1996;9:563- 84.

112. Singh M, Suresh K, Ho LC, Ng GC, Yap EH. Elucidation of the life cycle of the intestinal protozoan Blastocystis hominis. Parasitol Res. 1995;81:446-50.

113. Tan KS. Blastocystis in humans and animals: new insights using modern methodologies. Vet Parasitol. 2004;126:121-44.

114. Leelayoova S, Rangsin R, Taamasri P, Naaglor T, Thathaisong U, Mungthin M. Evidence of waterborne transmission of Blastocystis hominis. Am J Trop Med Hyg. 2004;70:658-62.

115. Leelayoova S, Siripattanapipong S, Thathaisong U, Naaglor T, Taamasri P, Piyaraj P, Mungthin M. Drinking water: a possible source of Blastocystis spp. subtype 1 infection in schoolchildren of a rural community in central Thailand. Am J Trop Med Hyg. 2008;79:401-6.

116. Li LH, Zhou XN, Du ZW, Wang XZ, Wang LB, Jiang JY, et al. Molecular epidemiology of human Blastocystis in a village in Yunnan province, China. Parasitol Int. 2007;56:281-6.

117. Al-Binali AM, Bello CS, El-Shewy K, Abdulla SE. The prevalence of parasites in commonly used leafy vegetables in South Western, Saudi Arabia. Saudi Med J. 2006;27:613-6.

118. Nagel R, Cuttell L, Stensvold CR, Mills PC, Bielefeldt-Ohmann H, Traub RJ. Blastocystis subtypes in symptomatic and asymptomatic family members and pets and response to therapy. Intern Med J. 2012;42:1187-95.

119. Scanlan PD, Knight R, Song SJ, Ackermann G, Cotter PD. Prevalence and genetic diversity of Blastocystis in family units living in the United States. Infect Genet Evol. 2016;45:95-97.

120. Abe N, Nagoshi M, Takami K, Sawano Y, Yoshikawa H. A survey of Blastocystis sp. in livestock, pets, and zoo animals in Japan. Vet Parasitol. 2002;106:203-12.

121. Lee MG, Stenzel DJ. A survey of Blastocystis in domestic chickens. Parasitol Res. 1999;85:109-17.

122. Cian A, El Safadi D, Osman M, Moriniere R, Gantois N, Benamrouz-Vanneste S, et al. Molecular Epidemiology of Blastocystis sp. in Various Animal Groups from Two French Zoos and Evaluation of Potential Zoonotic Risk. PLoS One. 2017;12:e0169659.

123. Wang W, Cuttell L, Bielefeldt-Ohmann H, Inpankaew T, Owen H, Traub RJ. Diversity of Blastocystis subtypes in dogs in different geographical settings. Parasit Vectors. 2013;6:215.

50 124. Casero RD, Mongi F, Sánchez A, Ramírez JD. Blastocystis and urticaria: Examination of subtypes and morphotypes in an unusual clinical manifestation. Acta Trop. 2015;148:156- 61.

125. Ramírez JD, Sánchez A, Hernández C, Flórez C, Bernal MC, Giraldo JC, et al. Geographic distribution of human Blastocystis subtypes in South America. Infect Genet Evol. 2016;41:32-5.

126. Macchioni F, Segundo H, Gabrielli S, Totino V, Gonzales PR, Salazar E, et al. Dramatic decrease in prevalence of soil-transmitted helminths and new insights into intestinal protozoa in children living in the Chaco region, Bolivia. Am J Trop Med Hyg. 2015;92:794- 6.

127. Malheiros AF, Stensvold CR, Clark CG, Braga GB, Shaw JJ. Short report: Molecular characterization of Blastocystis obtained from members of the indigenous Tapirapé ethnic group from the Brazilian Amazon region, Brazil. Am J Trop Med Hyg. 2011;85:1050-3.

128. David ÉB, Guimarães S, de Oliveira AP, Goulart de Oliveira-Sequeira TC, Nogueira Bittencourt G, Moraes Nardi AR, et al. Molecular characterization of intestinal protozoa in two poor communities in the State of São Paulo, Brazil. Parasit Vectors. 2015;8:103.

129. Santín M, Gómez-Muñoz MT, Solano-Aguilar G, Fayer R. Development of a new PCR protocol to detect and subtype Blastocystis spp. from humans and animals. Parasitol Res. 2011;109:205-12.

130. Ramírez JD, Sánchez LV, Bautista DC, Corredor AF, Flórez AC, Stensvold CR. Blastocystis subtypes detected in humans and animals from Colombia. Infect Genet Evol. 2014;22:223- 8.

131. Helenbrook WD, Shields WM, Whipps CM. Characterization of Blastocystis species infection in humans and mantled howler monkeys, Alouatta palliata aequatorialis, living in close proximity to one another. Parasitol Res. 2015;114:2517-25.

132. Jimenez-Gonzalez DE, Martinez-Flores WA, Reyes-Gordillo J, Ramirez-Miranda ME, Arroyo-Escalante S, Romero-Valdovinos M, et al. Blastocystis infection is associated with irritable bowel syndrome in a Mexican patient population. Parasitol Res. 2012;110:1269- 75.

133. Villalobos G, Orozco-Mosqueda GE, Lopez-Perez M, Lopez-Escamilla E, Córdoba-Aguilar A, Rangel-Gamboa L, et al. Suitability of internal transcribed spacers (ITS) as markers for the population genetic structure of Blastocystis spp. Parasit Vectors. 2014;7:461.

134. Vargas-Sanchez GB, Romero-Valdovinos M, Ramirez-Guerrero C, Vargas-Hernandez I, Ramirez-Miranda ME, Martinez-Ocaña J, et al. Blastocystis Isolates from Patients with Irritable Bowel Syndrome and from Asymptomatic Carriers Exhibit Similar Parasitological Loads, but Significantly Different Generation Times and Genetic Variability across Multiple Subtypes. PLoS One. 2015;10:e0124006.

135. Villegas-Gómez I, Martínez-Hernández F, Urrea-Quezada A, González-Díaz M, Durazo M, Hernández J, et al. Comparison of the genetic variability of Blastocystis subtypes between human carriers from two contrasting climatic regions of México. Infect Genet Evol. 2016;44:334-40.

136. Jones MS, Whipps CM, Ganac RD, Hudson NR, Boorom K. Association of Blastocystis subtype 3 and 1 with patients from an Oregon community presenting with chronic gastrointestinal illness. Parasitol Res. 2009;104:341-5.

137. Stensvold R, Brillowska-Dabrowska A, Nielsen HV, Arendrup MC. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J Parasitol. 2006;92:1081-7.

51 138. Stensvold CR, Arendrup MC, Jespersgaard C, Mølbak K, Nielsen HV. Detecting Blastocystis using parasitologic and DNA-based methods: a comparative study. Diagn Microbiol Infect Dis. 2007;59:303-7.

139. Rene BA, Stensvold CR, Badsberg JH, Nielsen HV. Subtype analysis of Blastocystis isolates from Blastocystis cyst excreting patients. Am J Trop Med Hyg. 2009;80:588-92.

140. Stensvold CR, Lewis HC, Hammerum AM, Porsbo LJ, Nielsen SS, Olsen KE, et al. Blastocystis: unravelling potential risk factors and clinical significance of a common but neglected parasite. Epidemiol Infect. 2009;137:1655-63.

141. Stensvold CR, Christiansen DB, Olsen KE, Nielsen HV. Blastocystis sp. subtype 4 is common in Danish Blastocystis-positive patients presenting with acute diarrhea. Am J Trop Med Hyg. 2011;84:883-5.

142. Stensvold CR, Nielsen SD, Badsberg JH, Engberg J, Friis-Møller N, Nielsen SS, et al. The prevalence and clinical significance of intestinal parasites in HIV-infected patients in Denmark. Scand J Infect Dis. 2011;43:129-35.

143. Andersen LO, Bonde I, Nielsen HB, Stensvold CR. A retrospective metagenomics approach to studying Blastocystis. FEMS Microbiol Ecol. 2015;91:fiv072.

144. Souppart L, Sanciu G, Cian A, Wawrzyniak I, Delbac F, et al. Molecular epidemiology of human Blastocystis isolates in France. Parasitol Res. 2009;105:413-21.

145. Poirier P, Wawrzyniak I, Albert A, El Alaoui H, Delbac F, Livrelli V. Development and evaluation of a real-time PCR assay for detection and quantification of Blastocystis parasites in human stool samples: prospective study of patients with hematological malignancies. J Clin Microbiol. 2011;49:975-83.

146. El Safadi D, Cian A, Nourrisson C, Pereira B, Morelle C, Bastien P, et al. Prevalence, risk factors for infection and subtype distribution of the intestinal parasite Blastocystis sp. from a large-scale multi-center study in France. BMC Infect Dis. 2016;16:451.

147. Yoshikawa H, Wu Z, Kimata I, Iseki M, Ali IK, Hossain MB, et al. Polymerase chain reaction-based genotype classification among human Blastocystis hominis populations isolated from different countries. Parasitol Res. 2004;92:22-9.

148. Menounos PG, Spanakos G, Tegos N, Vassalos CM, Papadopoulou C, Vakalis NC. Direct detection of Blastocystis sp. in human faecal samples and subtype assignment using single strand conformational polymorphism and sequencing. Mol Cell Probes. 2008;22:24-9.

149. Scanlan PD, Marchesi JR. Micro-eukaryotic diversity of the human distal gut microbiota: qualitative assessment using culture-dependent and -independent analysis of faeces. ISME J. 2008;2:1183-93.

150. Scanlan PD, Stensvold CR, Cotter PD. Development and Application of a Blastocystis Subtype-Specific PCR Assay Reveals that Mixed-Subtype Infections Are Common in a Healthy Human Population. Appl Environ Microbiol. 2015;81:4071-6.

151. Meloni D, Sanciu G, Poirier P, El Alaoui H, Chabé M, Delhaes L, et al. Molecular subtyping of Blastocystis sp. isolates from symptomatic patients in Italy. Parasitol Res. 2011;109:613- 9.

152. Mattiucci S, Crisafi B, Gabrielli S, Paoletti M, Cancrini G. Molecular epidemiology and genetic diversity of Blastocystis infection in humans in Italy. Epidemiol Infect. 2016;144:635-46.

153. Bart A, Wentink-Bonnema EM, Gilis H, Verhaar N, Wassenaar CJ, van Vugt M, et al. Diagnosis and subtype analysis of Blastocystis sp. in 442 patients in a hospital setting in the Netherlands. BMC Infect Dis. 2013;13:389.

52 154. Domínguez-Márquez MV, Guna R, Muñoz C, Gómez-Muñoz MT, Borrás R. High prevalence of subtype 4 among isolates of Blastocystis hominis from symptomatic patients of a health district of Valencia (Spain). Parasitol Res. 2009;105:949-55.

155. Alfellani MA, Stensvold CR, Vidal-Lapiedra A, Onuoha ES, Fagbenro-Beyioku AF, Clark CG. Variable geographic distribution of Blastocystis subtypes and its potential implications. Acta Trop. 2013;126:11-8.

156. Hussein EM, Hussein, AM, Eida MM, Atwa MM. Pathophysiological variability of different genotypes of human Blastocystis hominis Egyptian isolates in experimentally infected rats. Parasitol Res. 2008;102:853-60.

157. Souppart L, Moussa H, Cian A, Sanciu G, Poirier P, El Alaoui H, et al. Subtype analysis of Blastocystis isolates from symptomatic patients in Egypt. Parasitol Res. 2010;106:505–11.

158. Fouad SA, Basyoni MM, Fahmy RA, Kobaisi MH. The pathogenic role of different Blastocystis hominis genotypes isolated from patients with irritable bowel syndrome. Arab J Gastroenterol. 2011;12:194-200.

159. El Deeb HK, Khodeer S. Blastocystis spp.: frequency and subtype distribution in iron deficiency anemic versus non-anemic subjects from Egypt. J Parasitol. 2013;99:599-602.

160. Abdulsalam AM, Ithoi I, Al-Mekhlafi HM, Al-Mekhlafi AM, Ahmed A, Surin J. Subtype distribution of Blastocystis isolates in Sebha, Libya. PLoS One. 2013;8:e84372.

161. Poulsen CS, Efunshile AM, Nelson JA, Stensvold CR. Epidemiological Aspects of Blastocystis Colonization in Children in Ilero, Nigeria. Am J Trop Med Hyg. 2016;95:175- 9.

162. El Safadi D, Gaayeb L, Meloni D, Cian A, Poirier P, Wawrzyniak I, et al. Children of Senegal River Basin show the highest prevalence of Blastocystis sp. ever observed worldwide. BMC Infect Dis. 2014;14:164.

163. Petrášová J, Uzlíková M, Kostka M, Petrželková KJ, Huffman MA, Modrý D. Diversity and host specificity of Blastocystis in syntopic primates on Rubondo Island, Tanzania. Int J Parasitol. 2011;41:1113-20.

164. Pandey PK, Verma P, Marathe N, Shetty S, Bavdekar A, Patole MS, et al. Prevalence and subtype analysis of Blastocystis in healthy Indian individuals. Infect Genet Evol. 2015;31:296-9.

165. Das R, Khalil S, Mirdha BR, Makharia GK, Dattagupta S, Chaudhry R. Molecular Characterization and Subtyping of Blastocystis Species in Irritable Bowel Syndrome Patients from North India. PLoS One. 2016;11:e0147055.

166. Motazedian H, Ghasemi H, Sadjjadi SM. Genomic diversity of Blastocystis hominis from patients in southern Iran. Ann Trop Med Parasitol. 2008;102:85-8.

167. Moosavi A, Haghighi A, Mojarad EN, Zayeri F, Alebouyeh M, Khazan H, et al. Genetic variability of Blastocystis sp. isolated from symptomatic and asymptomatic individuals in Iran. Parasitol Res. 2012;111:2311-5.

168. Badparva E, Sadraee J, Kheirandish F, Frouzandeh M. Genetic diversity of human Blastocystis isolates in Khorramabad, central Iran. Iran J Parasitol. 2014;9:44-9.

169. Khoshnood S, Rafiei A, Saki J, Alizadeh K. Prevalence and genotype characterization of Blastocystis hominis among the Baghmalek people in Southwestern Iran in 2013 - 2014. Jundishapur J Microbiol. 2015;8:e23930.

53 170. Azizian M, Basati G, Abangah G, Mahmoudi MR, Mirzaei A. Contribution of Blastocystishominis subtypes and associated inflammatory factors in development of irritable bowel syndrome. Parasitol Res. 2016;11:2003-9.

171. Khademvatan S, Masjedizadeh R, Rahim F, Mahbodfar H, Salehi R, Yousefi-Razin E, Foroutan M. Blastocystis and irritable bowel syndrome: Frequency and subtypes from Iranian patients. Parasitol Int. 2017;66:142-5.

172. El Safadi D, Meloni D, Poirier P, Osman M, Cian A, Gaayeb L, et al. Molecular epidemiology of Blastocystis in Lebanon and correlation between subtype 1 and gastrointestinal symptoms. Am J Trop Med Hyg. 2013;88:1203-6.

173. Osman M, El Safadi D, Cian A, Benamrouz S, Nourrisson C, Poirier P, et al. Prevalence and Risk Factors for Intestinal Protozoan Infections with Cryptosporidium, Giardia, Blastocystis and Dientamoeba among Schoolchildren in Tripoli, Lebanon. PLoS Negl Trop Dis. 2016;10:e0004496.

174. Yoshikawa H, Wu Z, Pandey K, Pandey BD, Sherchand JB, Yanagi T, Kanbara H. Molecular characterization of Blastocystis isolates from children and rhesus monkeys in Kathmandu, Nepal. Vet Parasitol. 2009;160:295-300.

175. Lee IL, Tan TC, Tan PC, Nanthiney DR, Biraj MK, Surendra KM, Suresh KG. Predominance of Blastocystis sp. subtype 4 in rural communities, Nepal. Parasitol Res. 2012;110:1553-62.

176. Yakoob J, Jafri W, Beg MA, Abbas Z, Naz S, Islam M, Khan R. Irritable bowel syndrome: is it associated with genotypes of Blastocystis hominis. Parasitol Res. 2010;106:1033-8.

177. Abu-Madi M, Aly M, Behnke JM, Clark CG, Balkhy H. The distribution of Blastocystis subtypes in isolates from Qatar. Parasit Vectors. 2015;8:465.

178. Özyurt M, Kurt O, Mølbak K, Nielsen HV, Haznedaroglu T, Stensvold CR. Molecular epidemiology of Blastocystis infections in Turkey. Parasitol Int. 2008;57:300-6.

179. Dogruman-Al F, Dagci H, Yoshikawa H, Kurt O, Demirel M. A possible link between subtype 2 and asymptomatic infections of Blastocystis hominis. Parasitol Res. 2008;103:685-9.

180. Eroglu F, Genc A, Elgun G, Koltas IS. Identification of Blastocystis hominis isolates from asymptomatic and symptomatic patients by PCR. Parasitol Res. 2009;105:1589-92.

181. Dogruman-Al F, Kustimur S, Yoshikawa H, Tuncer C, Simsek Z, Tanyuksel M, et al. Blastocystis subtypes in irritable bowel syndrome and inflammatory bowel disease in Ankara, Turkey. Mem Inst Oswaldo Cruz. 2009;104:724-7.

182. Dogruman-Al F, Yoshikawa H, Kustimur S, Balaban N. PCR-based subtyping of Blastocystis isolates from symptomatic and asymptomatic individuals in a major hospital in Ankara, Turkey. Parasitol Res. 2009;106:263-8.

183. Eroglu F, Koltas IS. Evaluation of the transmission mode of B. hominis by using PCR method. Parasitol Res. 2010;107:841-5

184. Dagci H, Kurt Ö, Demirel M, Mandiracioglu A, Aydemir S, Saz U, et al. Epidemiological and diagnostic features of Blastocystis infection in symptomatic patients in Izmir province, Turkey. Iran J Parasitol. 2014;9:519-29.

185. Koltas IS, Eroglu F. Subtype analysis of Blastocystis isolates using SSU rRNA-DNA sequencing in rural and urban population in southern Turkey. Exp Parasitol. 2016;170:247-51.

54 186. Dogan N, Aydin M, Tuzemen NU, Dinleyici EC, Oguz I, Dogruman-Al F. Subtype distribution of Blastocystis spp. isolated from children in Eskisehir, Turkey. Parasitol Int. 2017;66:948-51.

187. Coskun A, Malatyali E, Ertabaklar H, Yasar MB, Karaoglu AO, Ertug S. Blastocystis in ulcerative colitis patients: Genetic diversity and analysis of laboratory findings. Asian Pac J Trop Med. 2016;9:916-9.

188. AbuOdeh R, Ezzedine S, Samie A, Stensvold CR, ElBakri A. Prevalence and subtype distribution of Blastocystis in healthy individuals in Sharjah, United Arab Emirates. Infect Genet Evol. 2016;37:158-62.

189. Wang W, Owen H, Traub RJ, Cuttell L, Inpankaew T, Bielefeldt-Ohmann H. Molecular epidemiology of Blastocystis in pigs and their in-contact humans in Southeast Queensland, Australia, and Cambodia. Vet Parasitol. 2014;203:264-9.

190. Yan Y, Su S, Lai R, Liao H, Ye J, Li X, Luo X, Chen G. Genetic variability of Blastocystis hominis isolates in China. Parasitol Res. 2006;99:597-601.

191. Li LH, Zhang XP, Lv S, Zhang L, Yoshikawa H, Wu Z, et al. Cross-sectional surveys and subtype classification of human Blastocystis isolates from four epidemiological settings in China. Parasitol Res. 2007;102:83-90.

192. Yoshikawa H, Tokoro M, Nagamoto T, Arayama S, Asih PB, Rozi IE, Syafruddin D. Molecular survey of Blastocystis sp. from humans and associated animals in an Indonesian community with poor hygiene. Parasitol Int. 2016;65:780-4.

193. Tan TC, Suresh KG, Smith HV. Phenotypic and genotypic characterisation of Blastocystis hominis isolates implicates subtype 3 as a subtype with pathogenic potential. Parasitol Res. 2008;104:85-93.

194. Tan TC, Ong SC, Suresh KG. Genetic variability of Blastocystis sp. isolates obtained from cancer and HIV/AIDS patients. Parasitol Res. 2009;105:1283-6.

195. Rivera WL, Tan MA. Molecular characterization of Blastocystis isolates in the Philippines by riboprinting. Parasitol Res. 2005;96:253-7.

196. Adao DE, Dela Serna A, Belleza , Bolo NR, Rivera WL. Subtype analysis of Blastocystis sp. isolates from asymptomatic individuals in an urban community in the Philippines. Ann Parasitol. 2016;62:193–200.

197. Belleza ML, Reyes JC, Tongol-Rivera PN, Rivera WL. Subtype analysis of Blastocystis sp. isolates from human and canine hosts in an urban community in the Philippines. Parasitol Int. 2016;65:291-4.

198. Wong KH, Ng GC, Lin RT, Yoshikawa H, Taylor MB, Tan KS. Predominance of subtype 3 among Blastocystis isolates from a major hospital in Singapore. Parasitol Res. 2008;102:663-70.

199. Thathaisong U, Worapong J, Mungthin M, Tan-Ariya P, Viputtigul K, Sudatis A, et al. Blastocystis isolates from a pig and a horse are closely related to Blastocystis hominis. J Clin Microbiol. 2003;41:967-75.

200. Jantermtor S, Pinlaor P, Sawadpanich K, Pinlaor S, Sangka A, Wilailuckana C, Wongsena W, Yoshikawa H. Subtype identification of Blastocystis spp. isolated from patients in a major hospital in northeastern Thailand. Parasitol Res. 2013;112:1781-6.

201. Thathaisong U, Siripattanapipong S, Mungthin M, Pipatsatitpong D, Tan-ariya P, Naaglor T, Leelayoova S. Identification of Blastocystis subtype 1 variants in the Home for Girls, Bangkok, Thailand. Am J Trop Med Hyg. 2013;88:352-8.

55 202. Sanpool O, Laoraksawong P, Janwan P, Intapan PM, Sawanyawisuth K, Thanchomnang T, Changtrakul Y, Maleewong W. Genetic subtypes of Blastocystis isolated from Thai hospitalized patients in Northeastern Thailand. Southeast Asian J Trop Med Public Health. 2015;46:184-90.

203. Popruk S, Udonsom R, Koompapong K, Mahittikorn A, Kusolsuk T, Ruangsittichai J, Palasuwan A. Subtype distribution of Blastocystis in Thai-Myanmar border, Thailand. Korean J Parasitol. 2015;53:13-9.

204. Palasuwan A, Palasuwan D, Mahittikorn A, Chiabchalard R, Combes V, Popruk S. Subtype Distribution of Blastocystis in Communities along the Chao Phraya River, Thailand. Korean J Parasitol. 2016;54:455-60.

205. Roberts T, Stark D, Harkness J, Ellis J. Subtype distribution of Blastocystis isolates identified in a Sydney population and pathogenic potential of Blastocystis. Eur J Clin Microbiol Infect Dis. 2013;32:335-43.

206. Levy Y, George J, Shoenfeld Y. Severe Blastocystis hominis in an elderly man. J Infect. 1996;33:57-9.

207. Stensvold CR, Arendrup MC, Nielsen HV, Bada A, Thorsen S. Symptomatic infection with Blastocystis sp. subtype 8 successfully treated with trimethoprim-sulfamethoxazole. Ann Trop Med Parasitol. 2008;102:271-4.

208. Vogelberg C, Stensvold CR, Monecke S, Ditzen A, Stopsack K, Heinrich-Gräfe U, Pöhlmann C. Blastocystis sp. subtype 2 detection during recurrence of gastrointestinal and urticarial symptoms. Parasitol Int. 2010;59:469-71.

209. Brandonisio O, Maggi P, Panaro MA, Lisi S, Andriola A, Acquafredda A, Angarano G. Intestinal protozoa in HIV-infected patients in Apulia, South Italy. Epidemiol Infect. 1999;123:457-62.

210. Cirioni O, Giacometti A, Drenaggi D, Ancarani F, Scalise G. Prevalence and clinical relevance of Blastocystis hominis in diverse patient cohorts. Eur J Epidemiol. 1999;15:389-93.

211. Leder K, Hellard ME, Sinclair MI, Fairley CK, Wolfe R. No correlation between clinical symptoms and Blastocystis hominis in immunocompetent individuals. J Gastroenterol Hepatol. 2005;20:1390-4.

212. Shlim DR, Hoge CW, Rajah R, Rabold JG, Echeverria P. Is Blastocystis hominis a cause of diarrhea in travelers? A prospective controlled study in Nepal. Clin Infect Dis. 1995;21:97- 101.

213. Giacometti A, Cirioni O, Fiorentini A, Fortuna M, Scalise G. Irritable bowel syndrome in patients with Blastocystis hominis infection. Eur J Clin Microbiol Infect Dis. 1999;18:436- 9.

214. Yakoob J, Jafri W, Jafri N, Khan R, Islam M, Beg MA, Zaman V. Irritable bowel syndrome: in search of an etiology: role of Blastocystis hominis. Am J Trop Med Hyg. 2004;70:383-5.

215. Ragavan ND, Kumar S, Chye TT, Mahadeva S, Shiaw-Hooi H. Blastocystis sp. in Irritable Bowel Syndrome (IBS)--Detection in Stool Aspirates during Colonoscopy. PLoS One. 2015;10:e0121173.

216. Tungtrongchitr A, Manatsathit S, Kositchaiwat C, Ongrotchanakun J, Munkong N, Chinabutr P, Leelakusolvong S, Chaicumpa W. Blastocystis hominis infection in irritable bowel syndrome patients. Southeast Asian J Trop Med Public Health. 2004;35:705-10.

56 217. Surangsrirat S, Thamrongwittawatpong L, Piyaniran W, Naaglor T, Khoprasert C, Taamasri P, et al. Assessment of the association between Blastocystis infection and irritable bowel syndrome. J Med Assoc Thai. 2010;93 Suppl 6:S119-24.

218. Krogsgaard LR, Engsbro AL, Stensvold CR, Nielsen HV, Bytzer P. The prevalence of intestinal parasites is not greater among individuals with irritable bowel syndrome: a population-based case-control study. Clin Gastroenterol Hepatol. 2015;13:507-13.

219. Puthia MK, Vaithilingam A, Lu J, Tan KS. Degradation of human secretory immunoglobulin A by Blastocystis. Parasitol Res. 2005;97:386-9.

220. Puthia MK, Sio SW, Lu J, Tan KS. Blastocystis ratti induces contact-independent apoptosis, F-actin rearrangement, and barrier function disruption in IEC-6 cells. Infect Immun. 2006;74:4114-23.

221. Puthia MK, Lu J, Tan KS. Blastocystis ratti contains cysteine proteases that mediate interleukin-8 response from human intestinal epithelial cells in an NF-kappaB-dependent manner. Eukaryot Cell. 2008;7:435-43.

222. Mirza H, Tan KS. Blastocystis exhibits inter- and intra-subtype variation in cysteine protease activity. Parasitol Res. 2009;104:355-61.

223. Nourrisson C, Wawrzyniak I, Cian A, Livrelli V, Viscogliosi E, Delbac F, Poirier P. On Blastocystis secreted cysteine proteases: a legumain-activated cathepsin B increases paracellular permeability of intestinal Caco-2 cell monolayers. Parasitology. 2016;143:1713-22.

224. Li J, Deng T, Li X, Cao G, Li X, Yan Y. A rat model to study Blastocytis subtype 1 infections.Parasitol Res. 2013;112:3537-41.

225. Leelayoova S, Taamasri P, Rangsin R, Naaglor T, Thathaisong U, Mungthin M. In-vitro cultivation: a sensitive method for detecting Blastocystis hominis. Ann Trop Med Parasitol. 2002;96:803-7.

226. Termmathurapoj S, Leelayoova S, Aimpun P, Thathaisong U, Nimmanon T, Taamasri P, Mungthin M. The usefulness of short-term in vitro cultivation for the detection and molecular study of Blastocystis hominis in stool specimens. Parasitol Res. 2004;93:445-7.

227. Stensvold CR, Ahmed UN, Andersen LO, Nielsen HV. Development and evaluation of a genus-specific, probe-based, internal-process-controlled real-time PCR assay for sensitive and specific detection of Blastocystis spp. J Clin Microbiol. 2012;50:1847-51.

228. Roberts T, Ellis J, Harkness J, Marriott D, Stark D. Treatment failure in patients with chronic Blastocystis infection. J Med Microbiol. 2014;63:252-7.

229. Coyle CM, Varughese J, Weiss LM, Tanowitz HB. Blastocystis: to treat or not to treat... Clin Infect Dis. 2012 Jan 1;54:105-10.

230. Cassano N, Scoppio BM, Loviglio MC, Vena GA. Remission of delayed pressure urticaria after eradication of Blastocystis hominis. Acta Derm Venereol. 2005;85:357-8.

231. Katsarou-Katsari A, Vassalos CM, Tzanetou K, Spanakos G, Papadopoulou C, Vakalis N. Acute urticaria associated with amoeboid forms of Blastocystis sp. subtype 3. Acta Derm Venereol. 2008;88:80-1.

232. Nigro L, Larocca L, Massarelli L, Patamia I, Minniti S, Palermo F, Cacopardo B. A placebo- controlled treatment trial of Blastocystis hominis infection with metronidazole. J Travel Med. 2003;10:128-30.

57 233. Armentia A, Méndez J, Gómez A, Sanchís E, Fernández A, de la Fuente R, Sánchez P. Urticaria by Blastocystis hominis. Successful treatment with paromomycin. Allergol Immunopathol (Madr). 1993;21:149-51.

234. Moghaddam DD, Ghadirian E, Azami M. Blastocystis hominis and the evaluation of efficacy of metronidazole and trimethoprim/sulfamethoxazole. Parasitol Res. 2005;96:273-5.

235. Mirza H, Teo JD, Upcroft J, Tan KS. A rapid, high-throughput viability assay for Blastocystis spp. reveals metronidazole resistance and extensive subtype-dependent variations in drug susceptibilities. Antimicrob Agents Chemother. 2011;55:637-48.

236. Haresh K, Suresh K, Khairul Anus A, Saminathan S. Isolate resistance of Blastocystis hominis to metronidazole. Trop Med Int Health. 1999;4:274-7.

237. Ok UZ, Girginkardeşler N, Balcioğlu C, Ertan P, Pirildar T, Kilimcioğlu AA. Effect of trimethoprim-sulfamethaxazole in Blastocystis hominis infection. Am J Gastroenterol. 1999;94:3245-7.

238. Rossignol JF, Kabil SM, Said M, Samir H, Younis AM. Effect of nitazoxanide in persistent diarrhea and enteritis associated with Blastocystis hominis. Clin Gastroenterol Hepatol. 2005;3:987-91.

239. Speich B, Marti H, Ame SM, Ali SM, Bogoch II, Utzinger J, et al. Prevalence of intestinal protozoa infection among school-aged children on Pemba Island, Tanzania, and effect of single-dose albendazole, nitazoxanide and albendazole-nitazoxanide. Parasit Vectors. 2013;6:3.

240. Valsecchi R, Leghissa P, Greco V. Cutaneous lesions in Blastocystis hominis infection. Acta Derm Venereol. 2004;84:322-3.

241. Kick G, Rueff F, Przybilla B. Palmoplantar pruritus subsiding after Blastocystis hominis eradication. Acta Derm Venereol. 2002;82:60.

242. Dinleyici EC, Eren M, Dogan N, Reyhanioglu S, Yargic ZA, Vandenplas Y. Clinical efficacy of Saccharomyces boulardii or metronidazole in symptomatic children with Blastocystis hominis infection. Parasitol Res. 2011;108:541-5.

243. Roberts T, Bush S, Ellis J, Harkness J, Stark D. In Vitro Antimicrobial Susceptibility Patterns of Blastocystis. Antimicrob Agents Chemother. 2015;59:4417-23.

244. Qvarnstrom Y, James C, Xayavong M, Holloway BP, Visvesvara GS, Sriram R, da Silva AJ. Comparison of real-time PCR protocols for differential laboratory diagnosis of amebiasis. J Clin Microbiol. 2005;43:5491-7.

245. Verweij JJ, Schinkel J, Laeijendecker D, van Rooyen MA, van Lieshout L, Polderman AM. Real-time PCR for the detection of Giardia lamblia. Mol Cell Probes. 2003;17:223-5.

246. Fontaine M, Guillot E. Development of a TaqMan quantitative PCR assay specific for Cryptosporidium parvum. FEMS Microbiol Lett. 2002;214:13-7.

247. Verweij JJ, Mulder B, Poell B, van Middelkoop D, Brienen EAT, van Lieshout L. Real-time PCR for the detection of Dientamoeba fragilis in fecal samples, Mol Cell Probes. 2007;21:400-4.

248. Vanni I, Cacciò SM, van Lith L, Lebbad M, Svärd SG, Pozio E, Tosini F. Detection of Giardia duodenalis assemblages A and B in human feces by simple, assemblage-specific PCR assays. PLoS Negl Trop Dis. 2012;6:e1776.

58 249. Johnson DW, Pieniazek NJ, Griffin DW, Misener L, Rose JB. Development of a PCR protocol for sensitive detection of Cryptosporidium oocysts in water samples. Appl Environ Microbiol. 1995;61:3849-55.

250. Lebbad M, Svärd SG. PCR differentiation of Entamoeba histolytica and Entamoeba dispar from patients with amoeba infection initially diagnosed by microscopy. Scand J Infect Dis. 2005;37:680-5.

251. Núñez YO, Fernández MA, Torres-Núñez D, Silva JA, Montano I, Maestre JL, Fonte L. Multiplex polymerase chain reaction amplification and differentiation of Entamoeba histolytica and Entamoeba dispar DNA from stool samples. Am J Trop Med Hyg. 2001;64:293-7.

252. Stensvold CR. Comparison of sequencing (barcode region) and sequence-tagged-site PCR for Blastocystis subtyping. J Clin Microbiol. 2013;51:190-4.

253. Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4:406-25.

254. Lebbad M, Petersson I, Karlsson L, Botero-Kleiven S, Andersson JO, Svenungsson B, Svärd SG. Multilocus genotyping of human Giardia isolates suggests limited zoonotic transmission and association between assemblage B and flatulence in children. PLoS Negl Trop Dis. 2011;5:e1262

255. Angelin M, Forsell J, Granlund M, Evengård B, Palmgren H, Johansson A. Risk factors for colonization with extended-spectrum beta-lactamase producing Enterobacteriaceae in healthcare students on clinical assignment abroad: A prospective study. Travel Med Infect Dis. 2015;13:223-9.

256. Bengtsson-Palme J, Angelin M, Huss M, Kjellqvist S, Kristiansson E, Palmgren H, et al. The Human Gut Microbiome as a Transporter of Antibiotic Resistance Genes between Continents. Antimicrob Agents Chemother. 2015;59:6551-60.

257. Bengtsson-Palme J, Hartmann M, Eriksson KM, Pal C, Thorell K, Larsson DG, Nilsson RH. METAXA2: improved identification and taxonomic classification of small and large subunit rRNA in metagenomic data. Mol Ecol Resour. 2015;15:1403-14.

258. Bengtsson-Palme J, Thorell K, Wurzbacher C, Sjöling Å, Nilsson, RH. Metaxa2 Diversity Tools: Easing microbial community analysis with Metaxa2. Ecological Informatics. 2016;33:45-50.

259. Benjamini Y, Hochberg Y. Controlling the false discovery rate: a practical and powerful approach to multiple testing. J Roy Stat Soc B Met. 1995;57:289-300.

260. Bhattacharya S, Bhattacharya A, Diamond LS, Soldo AT. The ribosomal DNA plasmids of Entamoeba. Parasitol Today. 1989;14:181-5.

261. van den Bijllaardt W, Overdevest IT, Buiting AG, Verweij JJ. Rapid clearance of Giardia lamblia DNA from the gut after successful treatment. Clin Microbiol Infect. 2014;20:O972- 4.

262. Sandhu GS, Kline BC, Stockman L, Roberts GD. Molecular probes for diagnosis of fungal infections. J Clin Microbiol. 1995 Nov;33:2913-9.

263. Svenungsson B, Lagergren A, Ekwall E, Evengård B, Hedlund KO, Kärnell A, et al. Enteropathogens in adult patients with diarrhea and healthy control subjects: a 1-year prospective study in a Swedish clinic for infectious diseases. Clin Infect Dis. 2000;30:770- 8.

59 264. Scanlan PD, Stensvold CR, Rajilić-Stojanović M, Heilig HG, De Vos WM, O'Toole PW, Cotter PD. The microbial eukaryote Blastocystis is a prevalent and diverse member of the healthy human gut microbiota. FEMS Microbiol Ecol. 2014;90:326-30.

265. Rossen NG, Bart A, Verhaar N, van Nood E, Kootte R, de Groot PF, et al. Low prevalence of Blastocystis sp. in active ulcerative colitis patients. Eur J Clin Microbiol Infect Dis. 2015;34:1039-44.

266. Al-Mekhlafi MS, Azlin M, Nor Aini U, Shaik A, Sa'iah A, Fatmah MS, et al. Giardiasis as a predictor of childhood malnutrition in Orang Asli children in Malaysia. Trans R Soc Trop Med Hyg. 2005;99:686-91.

267. Prado MS, Cairncross S, Strina A, Barreto ML, Oliveira-Assis AM, Rego S. Asymptomatic giardiasis and growth in young children; a longitudinal study in Salvador, Brazil. Parasitology. 2005;131:51-6.

268. Boeke CE, Mora-Plazas M, Forero Y, Villamor E. Intestinal protozoan infections in relation to nutritional status and gastrointestinal morbidity in Colombian school children. J Trop Pediatr. 2010;56:299-306.

269. Berkman DS, Lescano AG, Gilman RH, Lopez SL, Black MM. Effects of stunting, diarrhoeal disease, and parasitic infection during infancy on cognition in late childhood: a follow-up study. Lancet. 2002;359:564-71.

270. Ajjampur SS, Koshy B, Venkataramani M, Sarkar R, Joseph AA, Jacob KS, et al. Effect of cryptosporidial and giardial diarrhoea on social maturity, intelligence and physical growth in children in a semi-urban slum in south India. Ann Trop Paediatr. 2011;31:205-12.

271. Di Cristanziano V, Santoro M, Parisi F, Albonico M, Shaali MA, Di Cave D, Berrilli F. Genetic characterization of Giardia duodenalis by sequence analysis in humans and animals in Pemba Island, Tanzania. Parasitol Int. 2014:63;438-41.

272. Ignatius R, Gahutu JB, Klotz C, Steininger C, Shyirambere C, Lyng M, et al. High prevalence of Giardia duodenalis Assemblage B infection and association with underweight in Rwandan children. PLoS Negl Trop Dis. 2012;6:e1677.

273. Gascón J, Vargas M, Schellenberg D, Urassa H, Casals C, Kahigwa E, et al. Diarrhoea in children under 5 years of age from Ifakara, Tanzania: a case-control study. J Clin Microbiol. 2001;38:4459-62.

274. Kitahara K, Suzuki J. Sporolactobacillus nov. subgen. J Gen Appl Microbiol. 1963;9:59-71.

275. Yanagida F, Suzuki K, Kaneko T, Kozaki M, Komagata K. Morphological, biochemical, and physiological characteristics of spore-forming lactic acid bacteria. J Gen Appl Microbiol. 1987;33:33-45.

276. Chang YH, Jung MY, Park IS, Oh HM, Sporolactobacillus vineae sp. nov., a spore-forming lactic acid bacterium isolated from vineyard soil. Int J Syst Evol Microbiol. 2008;58:2316- 20.

277. Fujita R, Mochida K, Kato Y, Goto K. Sporolactobacillus putidus sp. nov., an endospore- forming lactic acid bacterium isolated from spoiled orange juice. Int J Syst Evol Microbiol. 2014;60:1499-503.

278. Lan QX, Chen J, Lin L, Ye XL, Yan QY, Huang JF, Liu CC, Yang GW. Sporolactobacillus pectinivorans sp. nov., an anaerobic bacterium isolated from spoiled jelly. Int J Syst Evol Microbiol. 2016;66:4323-8.

279. Huang HY, Huang SY, Chen PY, King VA, Lin YP, Tsen JH. Basic characteristics of Sporolactobacillus inulinus BCRC 14647 for potential probiotic properties. Curr Microbiol. 2007;54:396-404.

60 280. Pessione E. Lactic acid bacteria contribution to gut microbiota complexity: lights and shadows. Front Cell Infect Microbiol 2012;2:86.

281. Thao ML, Moran NA, Abbot P, Brennan EB, Burckhardt DH, Baumann P. Cospeciation of psyllids and their primary prokaryotic endosymbionts. Appl Environ Microbiol. 2000;66:2898-05.

282. Nakabachi A, Yamashita A, Toh H, Ishikawa H, Dunbar HE, Moran NA, Hattori M. The 160-kilobase genome of the bacterial endosymbiont Carsonella. Science. 2006;314:267.

283. Stenzel DJ, Boreham PF. Bacteria-like endosymbionts in Blastocystis sp. Int J Parasitol. 1994;24:147-9.

284. O'Brien Andersen L, Karim AB, Roager HM, Vigsnæs LK, Krogfelt KA, Licht TR, Stensvold CR. Associations between common intestinal parasites and bacteria in humans as revealed by qPCR. Eur J Clin Microbiol Infect Dis. 2016;35:1427-31.

285. De Filippo C, Cavalieri D, Di Paola M, Ramazzotti M, Poullet JB, Massart S, et al. Impact of diet in shaping gut microbiota revealed by a comparative study in children from Europe and rural Africa. Proc Natl Acad Sci U S A. 2010. 107, 14691-6.

286. Wu GD, Chen J, Hoffmann C, Bittinger K, Chen YY, Keilbaugh SA, et al. Linking long-term dietary patterns with gut microbial enterotypes. Science. 2011;334:105-8.

287. Audebert C, Even G, Cian A, The Blastocystis Investigation Group, Loywick A, Merlin S, et al. 2016. Colonization with the enteric protozoa Blastocystis is associated with increased diversity of human gut bacterial microbiota. Sci Rep. 2016;5:25255.

288. Bäckhed F, Fraser CM, Ringel Y, Sanders ME, Sartor RB, Sherman PM, Versalovic J, Young V, Finlay BB. Defining a healthy human gut microbiome: current concepts, future directions, and clinical applications. Cell Host Microbe. 2012;12:611-22.

289. Shafquat A, Joice R, Simmons SL, Huttenhower C. Functional and phylogenetic assembly of microbial communities in the human microbiome. Trends Microbiol. 2014;22:261-6.

290. Lloyd-Price J, Abu-Ali G, Huttenhower C. The healthy human microbiome. Genome Med. 2016;8:51.

291. Stensvold CR, Alfellani M, Clark CG. Levels of genetic diversity vary dramatically between Blastocystis subtypes. Infect Genet Evol. 2012;12:263-73.

292. United States Census Bureau. General Demographic Characteristics (2010 Census, DP-1). Downloaded from factfinder.census.gov on 09 January 2017.

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