Figures
Abstract
Background
Giardia duodenalis is a flagellate protozoan that parasitizes humans and several other mammals. Protozoan contamination has been regularly documented at important environmental sites, although most of these studies were performed at the species level. There is a lack of studies that correlate environmental contamination and clinical infections in the same region. The aim of this study is to evaluate the genetic diversity of a set of clinical and environmental samples and to use the obtained data to characterize the genetic profile of the distribution of G. duodenalis and the potential for zoonotic transmission in a metropolitan region of Brazil.
Methodology/Principal Findings
The genetic assemblages and subtypes of G. duodenalis isolates obtained from hospitals, a veterinary clinic, a day-care center and important environmental sites were determined via multilocus sequence-based genotyping using three unlinked gene loci. Cysts of Giardia were detected at all of the environmental sites. Mixed assemblages were detected in 25% of the total samples, and an elevated number of haplotypes was identified. The main haplotypes were shared among the groups, and new subtypes were identified at all loci. Ten multilocus genotypes were identified: 7 for assemblage A and 3 for assemblage B.
Conclusions/Significance
There is persistent G. duodenalis contamination at important environmental sites in the city. The identified mixed assemblages likely represent mixed infections, suggesting high endemicity of Giardia in these hosts. Most Giardia isolates obtained in this study displayed zoonotic potential. The high degree of genetic diversity in the isolates obtained from both clinical and environmental samples suggests that multiple sources of infection are likely responsible for the detected contamination events. The finding that many multilocus genotypes (MLGs) and haplotypes are shared by different groups suggests that these sources of infection may be related and indicates that there is a notable risk of human infection caused by Giardia in this region.
Citation: Durigan M, Abreu AG, Zucchi MI, Franco RMB, de Souza AP (2014) Genetic Diversity of Giardia duodenalis: Multilocus Genotyping Reveals Zoonotic Potential between Clinical and Environmental Sources in a Metropolitan Region of Brazil. PLoS ONE 9(12): e115489. https://doi.org/10.1371/journal.pone.0115489
Editor: Sanjai Kumar, Food and Drug Administration, United States of America
Received: March 19, 2014; Accepted: November 24, 2014; Published: December 23, 2014
Copyright: © 2014 Durigan et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: The authors confirm that all data underlying the findings are fully available without restriction. All sequence files are available from the GenBank database (accession number(s) JN116442–JN116504, KM495697-KM495727 and KF922892-KF923021).
Funding: This work was supported by the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) through a research grant and a research fellowship awarded to APS (number 577032/2008-9 and 304914/2010-0), by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES - BioComputacional Program) and by the Fundação de Amparo à Pesquisa do Estado de São Paulo (FAPESP) through a graduate fellowship to MD (MS 2008/52197-4 and DD 2011/50413-4). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Giardia duodenalis (syn. G. lamblia and G. intestinalis) is a flagellate unicellular protozoan that parasitizes both humans and several domestic and wild animal species. The Giardia genus consists of six species that have been characterized based on their morphology and trophozoite ultrastructure. The species identified to date are G. agilis, which parasitizes amphibians; G. ardeae and G. psittaci, which parasitize birds; G. microti and G. muris, which parasitize rodents; and G. duodenalis, which parasitizes mammals [1]–[3]. Giardiasis is one of the most common waterborne diseases in the world. The cysts are transmitted via the fecal-oral rote through contaminated water or food, by contact with infected people or animals or through sexual contact.
Approximately 1.2 million new cases of giardiasis occur annually in the United States alone [4], and 280 million people per year are estimated to present this infection globally [5]. The high prevalence of this infection in developed countries and the increasing frequency of outbreaks in day-care centers have led to the inclusion of giardiasis in the World Health Organization's (WHO) Neglected Disease Initiative [6], and Giardia is now considered a re-emerging infectious agent [7]. Over 1.7 million people die every year from poor hygiene, a lack of sanitary infrastructure and a lack of clean water or sewage treatment [8]. Diarrheal diseases, including giardiasis, account for 15% of all deaths in children younger than 5 years of age [9].
The available data on the prevalence of G. duodenalis in non-human hosts are primarily confined to dogs, cats and livestock. Although human infections are of primary interest, Giardia has been reported in dogs and cats worldwide [10]. Surveys addressing the presence of Giardia infections in dogs have reported a high prevalence [11], [12]; the prevalence in cats is also of great concern from both a clinical and public health perspective [13], [14]. Similarly, Giardia infection in livestock is common and may be an important cause of economic losses, because its prevalence can reach 100% in calves [15]–[17].
Outbreaks of human giardiasis are frequently caused by contamination of drinking water [18] and a high prevalence of Giardia has been detected in surface waters [19], [20]. However, few studies have assessed the levels of clinical or environmental contamination. Four densely urbanized regions of São Paulo State were found to present higher rates than are tolerated by the United States Environmental Protection Agency (USEPA) regarding the risk of infection. Within this scenario, the metropolitan region of Campinas exhibited the highest percentage of positivity for Giardia and the highest risk of infection among these regions [21]. The detection of Giardia in surface waters may provide important clues regarding the environmental epidemiology of the parasite [22], especially in developing countries, where water and sewage treatment rates remain low.
G. duodenalis shows a high degree of genetic diversity, and isolates of this species have been grouped into genetic assemblages [23]. Although studies investigating the genetic diversity of Giardia were initiated through allozymes [24] and restriction fragment length polymorphism (RFLP) analyses [25], the genetic assemblages of G. duodenalis are now mainly identified using PCR followed by sequencing of conserved genes such as glutamate dehydrogenase, triose phosphate isomerase, beta-giardin, elongation factor 1-alpha and small subunit ribosomal RNA (ssu-rRNA) [26]. Single nucleotide polymorphisms (SNPs) identified in amplified fragments of these genes have enabled the characterization of Giardia isolates in different genetic assemblages. G. duodenalis assemblages A and B show zoonotic potential and can parasitize many hosts, such as humans, dogs, cats, non-human primates, livestock, horses, pigs, deer, moose and ferrets. Cysts recovered from human samples have always been grouped into one of two major genetic assemblages (A and B) that have been described worldwide [24], [25], [27]–[30]. Molecular studies have revealed the existence of genetic structure within these major assemblages [31]. Approximately three-quarters of assemblage A sequences obtained from non-human hosts, such as livestock and companion animals, correspond to sub-assemblage AI. In humans, a similar proportion of assemblage A sequences have been genotyped as belonging to sub-assemblage AII [12], [32]. Assemblage B sub-assemblages BIII and BIV have been observed in cattle, horses, monkeys and dogs. In humans, the two sub-assemblages are observed at similar frequencies. Sub-assemblage AIII was first identified in wild hoofed stock and is restricted to animals. Thus, sub-assemblages AI, AII, BIII and BIV are potentially zoonotic [12], [32]–[34]. The other assemblages are commonly associated with specific hosts. For example, assemblages C and D have been identified in dogs, cats, coyotes and wolves; assemblage E has been found in livestock; assemblage F has been found in cats; and assemblage G has been found in rats [26], [31]. Recently, another assemblage (H) was identified in marine vertebrates as a sister clade to assemblage G based on an analysis of the gdh gene [35].
In the present study, Giardia spp. cysts obtained from multiple hosts and collected from different locations in urban areas of the metropolitan region of Campinas in the state of São Paulo, Brazil were genetically characterized based on multilocus analyses. The aim of this study is to determine the genetic diversity of a set of clinical and environmental samples and to use these data to assess the potential for zoonotic transmission and mixed infection and their possible impacts on public health.
Methods
Study area and design
Samples from multiple sites and hosts were collected to genetically characterize the cysts of the protozoan parasite Giardia spp. The Campinas metropolitan region in the state of São Paulo in southeastern Brazil consists of 20 cities that are home to nearly 3 million people. Campinas is the main city and includes three primary urban drainage basins that drain the majority of the water in this region. Environmental samples were collected from correlated water sources including rivers, urban streams, hospital sewage, a waste water treatment plant and the water abstraction site for the city from the Atibaia River. Clinical samples were obtained through partnerships with hospitals, a day-care center, a veterinary clinic and a farm, all of which provided positive samples (Fig. 1). Together, these partnerships allowed us to survey a greater number of parasitized hosts, thereby increasing the diversity of the sampled Giardia cysts. Because our collaborators provided positive samples, the focus of this study was not to present information about the prevalence of Giardia at the collection sites. The locations in the region where the environmental and clinical samples were collected share the same drainage basins and were strategically selected to aid in the identification of contamination events and the detection of relationships among samples of different origins. No clinical information was obtained for the isolates.
Brazil (a), state of São Paulo (b), metropolitan region of Campinas (c) and the collection sites of positive samples plotted on a map of the city of Campinas colored to indicate the various drainage basins (d). The assemblages identified in each group were plotted in the map close to the collection site. •Unicamp Hospital and hospital sewage. ⧫Proença stream, Serafim stream and day-care center. ▪Ouro Verde Hospital. ⧫Campinas water abstraction site. ★Anhumas River. ▴Samambaia waste water treatment plant.
Clinical samples
Giardia spp. cysts were obtained from the feces of humans, dogs, cats and a calf. The cysts from the human samples were obtained from children at a day-care center in the city of Campinas and from patients at two hospitals: the University of Campinas Clinic Hospital and the Ouro Verde Hospital both of which are located in Campinas. The isolates obtained from dogs and cats were provided by the Animaltec Veterinary Clinic in the city of Campinas. The calf sample was obtained from São Francisco Farm in Amparo, located 37 km northeast of Campinas in the state of São Paulo. The samples were labeled and grouped according to their origin. The hospital samples were labeled “HC samples”, the day-care samples were labeled “DC samples”, and the clinical samples from non-human hosts were labeled “VET samples”.
Ethics statement
The ethical aspects related to the acquisition of human clinical samples were approved by the ethics committee of the Faculdade de Ciências Médicas - Unicamp (251-2009). Written informed consent was obtained from legal guardians of all participants. No specific permissions were required for isolated Giardia cysts provided by our collaborators (VET and HC isolates). The Giardia cysts obtained from dogs and cats were provided by Edson Golçalves from the Animaltec Veterinary Clinic in the city of Campinas, with verbal consent of the owners, in accordance with the purposes of the research. The calf sample was obtained from São Francisco Farm in Amparo (22°42'04" S and 46°45'52" W) This sample was provided by Luciana Urbano do Santos, with consent of the owner, in accordance with the purposes of the research. Water samples were collected from hospital sewage at the University of Campinas Clinic Hospital, effluent from the Samambaia Waste Water Treatment Plant (SWWTP) (22°56′00″ S and 47°00′00″ W), a water abstraction site in Campinas (Atibaia River), the Proença urban stream (22° 53′ 16″ S and 47° 02′ 62″ W) and the Serafim urban stream (22° 53′ 20″ S and 47° 02′ 92″ W), which merge and form the Anhumas River, where another sample was collected (22° 52′ 80″ S and 47° 02′ 32″ W). No specific permissions were required for the environmental locations where the samples were collected. The field studies did not involve endangered or protected species.
Environmental samples
Water samples were collected from hospital sewage at the University of Campinas Clinic Hospital, effluent from the Samambaia Waste Water Treatment Plant (SWWTP), a water abstraction site in Campinas (Atibaia River), and the Proença and Serafim urban streams, which merge and form the Anhumas River, where another sample was collected. These samples were labeled “ENV samples”.
Parasitological techniques
A small aliquot obtained from the original suspension of cysts was re-examined for Giardia cysts using centrifugal flotation in saturated zinc sulfate [36] followed by microscopy. Following reconfirmation, the positive samples were purified on a sucrose gradient and concentrated. Antibiotics and a fungicide were subsequently added according to the original purification protocol [37].
The environmental samples were collected in bottles of different volumes, depending on the origin of the sample, and filtered (4 l/min), using a sterile mixed cellulose ester membrane with a diameter of 47 mm and a nominal porosity of 3 µm [19]. The samples were eluted by scraping the membrane with a smooth-edged plastic loop and rinsing the membrane with a 0.1% Tween 80 elution solution (RM method) [38]. The eluates were then centrifuged (1,050×g; 15 min), and the pellet was washed, centrifuged again and diluted in 1.5 ml of water from a 0.22 µm water filtration system (MilliQ, Millipore, Brazil). The only exception was the raw sewage sample obtained from sewage from the University of Campinas Clinic Hospital, which was concentrated via centrifugal sedimentation [39]. The pelleted samples were processed through immunomagnetic separation (IMS) using the Dynabead GC-Combo Kit followed by chemical dissociation according to the manufacturer's instructions (Dynal Biotech, Oslo, Norway). The positive samples were further confirmed using a direct immunofluorescence assay (DFA) according to the manufacturer's protocol. Briefly, 5 µl of each concentrated sample was fixed in a well slide with methanol, and fluorescein isothiocyanate-conjugated monoclonal antibodies specific for cell wall antigens from Merifluor (Cryptosporidium/Giardia Test Kit, Meridian Bioscience Inc., Cincinnati, OH, USA) were added. The cysts were identified based on their similar sizes, shapes and patterns/intensities of immunofluorescent staining using a Nikon (50i) microscope with a 450–490 nm excitation filter and a 520 nm barrier filter. The positive samples were stored in microcentrifuge tubes and frozen in liquid nitrogen to preserve their cellular integrity.
Molecular methods
DNA extraction.
DNA was extracted from clinical and environmental samples using the ZR Fungal/Bacterial DNA Kit (Zymo Research). Extraction was performed according to the original protocols provided by the manufacturer. Qualitative and quantitative estimation of each sample was performed via gel electrophoresis in a 1% agarose gel and with a NanoDrop 8000 Spectrophotometer (Thermo Scientific), respectively.
Molecular identification and genotyping of G. duodenalis assemblages.
All of the extracted samples were analyzed through the amplification of fragments of the beta-giardin gene (bg) to confirm the presence of Giardia. The reactions were performed using the same reagent concentrations and thermal cycling conditions previously described with the exception of the reaction volume of 25 µl [40]. A negative control consisting of the reaction mixture without DNA was included in each run. The reactions were performed in a C1000 Thermal Cycler (BioRad) and the amplified DNA fragments were detected through gel electrophoresis in a 3% agarose gel and visualized using the Gene Genius Bio Imaging System (Syngene) after staining with ethidium bromide.
Fragments consisting of 530 bp of the triose phosphate isomerase gene (tpi), 753 bp of the beta-giardin gene (bg) and 218 bp of the glutamate dehydrogenase gene (gdh) were individually amplified. These markers are single-copy genes and are unlinked in the G. duodenalis genome [41]. Each isolate was characterized using nested tpi PCR, nested gdh PCR and bg PCR [42]–[44], performed according to the original protocols, with the addition of 400 ng/ µl of bovine serum albumin (BSA) in the first reaction [45]. The VET samples were analyzed with C- and D- specific tpi primers [46]. Following the amplification reaction, the PCR products were purified using the QIAquick PCR Purification Kit (QIAGEN) and sequenced using the ABI Prism Big Dye Terminator Cycle Sequencing Kit, version 3.1 (Applied Biosystems). The sequencing reactions were analyzed on an ABI PRISM 3500 Genetic Analyser xL (Applied Biosystems) and all of the sequences were obtained through direct sequencing in both directions a minimum of four times. The sequences were edited using CLC Main Workbench v6.8.1 (CLC Bio.) and were aligned with reference sequences using ClustalX [47]. A set of isolates with sequences that presented double peaks was cloned into the PGEM T-Easy vector (Promega Corporation). Following the transfection of Escherichia coli XL1-blue, 50 recombinant clones were selected for DNA sequencing.
Phylogenetic reconstruction and analysis of genealogical relationships were implemented to characterize the sequences obtained in this study based on a comparison with previously characterized sequences. Reference bg, gdh, and tpi sequences from all of the major genetic assemblages of G. duodenalis were obtained from GenBank (Table S1 in S1 File). The genotyping of each isolate was performed at four different levels of resolution. Each isolate was assigned to a specific assemblage, followed by the characterization of sub-assemblages through phylogenetic analysis. The heterogeneous profiles identified within each sub-assemblage were described as subtypes. Subtypes from the same isolate obtained for different loci were combined to generate multilocus genotypes.
Accession numbers of the nucleotide sequences.
The accession numbers and the corresponding assemblages of the sequences obtained from the GenBank database are displayed in Table S1 in S1 File. The new sequences generated in this study have been deposited in GenBank under accession numbers JN116442-JN116504, KM495697-KM495727 and KF922892-KF923021. The accession numbers are also displayed in Tables S2, S3, S4 and S5 in S1 File.
Phylogenetic analysis and genealogical relationships among sequences
Phylogenetic analysis.
Neighbor-joining (NJ) and maximum likelihood (ML) analyses were performed using MEGA, version 5.05 [48]. The nucleotide substitution model that best fit the data was selected following analysis using jModelTest [49]. The analyses were conducted using the Tamura-Nei93 distance [50] with gamma correction, which assumes inequality in base frequencies and rate variation among sites. A bootstrap phylogeny test was performed with 10,000 replicates.
The Bayesian phylogenetic analysis was performed with MrBayes software version 3.1.2 [51] using the GTR model with gamma correction, which is the available model that is most similar to Tamura-Nei93. The starting trees were random, Markov chains were run for 6,000,000 iterations, and the trees were sampled every 500 iterations. Bayesian posterior probabilities were calculated using a Markov chain Monte Carlo sampling technique. Subtypes obtained from three loci were combined to define multilocus genotypes (MLGs). Isolates showing mixed assemblage profiles or overlapping nucleotides were not included in the multilocus analysis. The Bayesian inferred trees were visualized with TreeView X [52].
Estimation of genealogical relationships.
The aligned sequences were converted to the NEXUS format [53]. The genealogical relationships between sequences were estimated using TCS software [54], which collapses identical sequences into unique haplotypes and then generates a cladogram estimate. Missing data were not considered in the haplotype analyses. An absolute distance matrix was calculated for all pairwise comparisons of haplotypes. The probability of parsimony was calculated for pairwise differences until the probability exceeded 0.95. DNAsp software v5.10 [55] was used to generate an Arlequin data file. Arlequin software was employed to calculate haplotype diversity at the intra-population level [56].
Results
Detection of positive samples using parasitological methods
All of the surveyed environmental and clinical sites (Fig. 1) provided samples that were positive for Giardia spp.; in total, 91 positive samples were obtained. Among the human clinical samples, 51 positive samples were provided by hospitals (HC group) and 28 positive samples from children were provided by the day-care center (DC group). Six positive samples were obtained from non-human hosts (VET group): three from dogs (VET01, VET02 and VET03), two from cats (VET04 and VET05) and one from a calf (VET06). Among the environmental samples (ENV group), one positive sample was obtained from the SWWTP (ENV01), one from the water abstraction site of the Atibaia River (ENV02) and one from hospital sewage from the University of Campinas Clinic Hospital (ENV06). Two positive samples were identified from urban streams: one from the Proença stream (ENV03) and one from the Serafim stream (ENV04). These streams merge to form the Anhumas River, where another positive sample was identified (ENV05). In total, 87% (79/91) of the obtained Giardia cysts came from human clinical specimens, 6.5% (6/91) were from non-human specimens and 6.5% (6/91) were from environmental samples.
Molecular analyses
Molecular identification of G. duodenalis.
The results of the amplification of the fragments of the three genes were used to confirm the presence of Giardia in all of the samples. Consensus analysis of the complementary molecular methods (gdh, tpi, and both approaches for bg) confirmed 97% (88/91) of the samples to be positive. Amplification reactions targeting two different fragments of the bg gene [40] confirmed 51% (46/91) positivity. The amplification rates (followed by sequencing) of the three other markers used were 89% (81/91), 69% (63/91) and 49% (45/91) for the tpi, gdh and bg genes, respectively. Three samples (HC03, HC05 and HC26) were positive by DFA but repeatedly negative in all of the amplification reactions.
Molecular genotyping at the beta-giardin (bg) locus.
Among the 45 bg sequences generated, assemblage A was identified in 34 isolates: 15 from the HC group and 19 from the DC group. Nine assemblage B isolates were identified in the HC group. One assemblage D isolate and one assemblage E isolate were found in the VET group in samples from a dog (VET01) and a calf (VET 06), respectively. No sequence data were obtained for this marker in the ENV group. The distribution of each genetic assemblage within each source of contamination is presented in Table 1.
Molecular genotyping at the triose phosphate isomerase (tpi) locus.
Among the 81 tpi sequences obtained through the reaction with generic primers, assemblage A was identified in 20 HC, five DC and three VET isolates. Forty-seven isolates (21 from HC, 20 from DC, two from VET and four from ENV) were classified as belonging to assemblage B, which was the most prevalent assemblage at this locus. Five assemblage C isolates from the HC group and another one from the ENV group were identified. When tpi primers specific for assemblages C and D were used, sequencing revealed assemblage D in two isolates (VET01 and VET05) and assemblage C in an additional isolate (VET02), both from the VET group. Remarkably, these three samples had previously been characterized as belonging to different assemblages using the generic tpi primers.
Molecular genotyping at the glutamate dehydrogenase (gdh) locus.
At the gdh locus, five different assemblages were identified. Assemblage A was detected in 28 isolates (20 from HC, seven from DC and one from the ENV group). Assemblage B was found in 29 isolates (17 from HC, 11 from DC and one from the ENV group). Four isolates were characterized as belonging to assemblage D (three from VET and one from the ENV group). The isolate obtained from the calf was genotyped as belonging to assemblage E (VET06), and one isolate obtained from a dog was characterized as assemblage C (VET03). Table S6 in S1 File presents the comparison of each source of contamination and its distribution within each genetic assemblage.
Combined genotyping results at the assemblage and sub-assemblage levels.
A total of 192 sequences were generated in this study: 63 for the gdh gene, 45 for the bg gene, 81 for the tpi gene and three additional sequences through assemblage-specific tpi PCR. Sequence data were obtained for three genes from 37 samples, for two genes from 29 samples and for one gene from 22 samples. In the sequenced hospital samples (n = 48), assemblages A (42%), B (42%) and C (6%) were identified. Contrasting assemblage results between markers were found in five isolates. All assemblage A isolates from the HC group corresponded to sub-assemblage AII. Among the assemblage B isolates from this group, only one (HC-33) was characterized as BIII, whereas the remainder were BIV. Similar results were observed in the DC group, in which all assemblage A isolates were characterized as AII. Five isolates were genotyped as BIV and four as BIII. Twelve samples presented different assemblage results between markers. Many genetic assemblages were found in the VET group, in which most of the isolates from dogs and cats and that from the calf were characterized as belonging to host-adapted assemblages (C and D for dogs and cats and E for the calf). However, the sequences generated using the generic tpi primers were characterized as belonging to assemblages with zoonotic potential (S1 Fig.). Most of the isolates from the ENV group exhibited assemblages with zoonotic potential even though assemblages C and D were identified at the water abstraction point of the city (ENV02) and in the Anhumas River (ENV05), respectively. The genotyping results for each sample are presented in Tables S2, S3, S4, and S5 in S1 File. The identified assemblages were plotted on the physical map close to their respective collection sites (Fig. 1).
Mixed assemblages and heterogeneous sequencing profiles.
All of the surveyed groups presented at least one sample in which the assignment to a specific assemblage was inconsistent when two or more different loci were compared. Mixed assemblages in an individual isolate were identified in 26% (23/88) of all sequenced samples. When considering only isolates that were sequenced with at least two different markers or were cloned, the proportion of mixed assemblages increased to 34% (23/66). In the HC group, 12% (6/48) of the isolates were identified as showing this pattern. Two human isolates presented both assemblages A and C, and another three isolates presented mixtures of A and B. One isolate (HC-07) exhibited assemblages BIII and BIV. Within the DC group, 39% (11/28) of the isolates presented mixed assemblages, all of which involved assemblages A and B. The highest prevalence of mixed assemblages was found in the VET group, in which 83% (5/6) of the samples were characterized as displaying different assemblages. This prevalence increased to 100% when considering only the samples genotyped with at least two markers. In two cases, more than two assemblages were observed in individual VET samples (VET02 and VET04). In the ENV group, only isolate ENV-05 presented a mixed assemblage (BIV and D), even though most of the samples (4/6) were successfully sequenced with only one marker. Of the 22 isolates with mixed assemblages identified, 64% (14/22) involved assemblages A and B. Three isolates presented assemblages B and D, two presented A and C, and one presented A and E. Table 2 provides the genetic assemblages of each isolate in which mixed assemblages were identified.
Overlapping nucleotides in the chromatograms were verified in a considerable number of sequences. Thirty-seven percent (22/63) of the gdh sequences presented double peaks, whereas 27% (11/45) of the bg sequences and 26% (21/84) of the tpi sequences presented this pattern. In all of the assemblages, at least one isolate exhibited double peaks in the chromatogram. Most of the samples in which double peaks were observed were characterized as belonging to assemblage B. To assure reproducibility, overlapping nucleotides were only considered when the same pattern was observed in different sequencing reactions for the same sample. A complete list with all isolates that presented double peaks is displayed in Table S7 in S1 File.
Following the cloning of fragments with double peaks, additional sequences were detected: three new sequences were identified from the VET04 sample (one BIV, one BIII and one C) and three from the VET02 sample (two C and one AI). Three different subtypes were identified from the HC43 sample (all BIV). The sequences from HC51(gdh), VET06(tpi), DC04(tpi), DC05(tpi) and ENV06(gdh) presented only one haplotype phase.
Subtypes.
Subtypes could be determined only when double peaks were not identified at any position. At the bg locus, 33 of the 45 (73%) sequenced isolates exhibited no double peaks. These 33 isolates corresponded to 13 subtypes, including nine new subtypes and four previously reported subtypes. The novel subtypes were represented by one isolate each (Table S8 in S1 File). At the gdh locus, 40 of 63 (63%) isolates presented no overlapping nucleotides in the chromatogram. Eight different subtypes were observed, two of which corresponded to novel subtypes. The most common subtype was found in 21 isolates (Table S9 in S1 File). Among the 84 sequences obtained at the tpi locus, 62/84 (74%) exhibited no double peaks. Thirty different subtypes were detected, 21 of which were new. The most common subtype was observed in 17 isolates (Table S10 in S1 File).
Genealogical relationships.
A similar proportion of haplotypes was found when different molecular markers were compared. As the focus of this analysis was not the isolates but the haplotypes, isolates with overlapping nucleotides were included. In total, 20, 49 and 24 different haplotypes were identified for the bg, tpi and gdh genes, respectively (Table 3).
Considering all of the examined molecular markers, assemblage B was the most common, with 50 haplotypes, followed by assemblage A, with 25. Assemblages C, D and E presented 11, five and two different haplotypes, respectively. Most haplotypes from the tpi and gdh genes were characterized as belonging to assemblage B. At the bg locus, there were similar proportions of A and B. The relative frequencies of each source within the main haplotypes are presented in Fig. 2.
The relative frequencies of the main haplotypes from the day-care center (green), hospital clinical (dark blue), veterinary (yellow) and environmental (red) samples generated from a statistical parsimony network. The sizes of the circles are proportional to the haplotype frequency. The lines connecting each haplotype represent one mutation, and small white circles represent inferred haplotypes that were not actually observed. Haplotypes that are not connected to others exhibited more steps (or mutations) than the connection limit (95% confidence).
Many haplotypes were found in the four groups. The HC group exhibited 48 different haplotypes, most of which were from assemblage B isolates. A comparable result was observed in the DC group, in which 31 different haplotypes were found (61% from assemblage B), and in the ENV group, in which 62.5% (5/8) of the haplotypes were from assemblage B. In contrast, the VET group presented a balanced distribution of haplotypes in many assemblages (Table 4). Haplotype variation was detected in almost all of the environmental samples. A complete list of haplotypes is provided in Tables S11, S12 and S13 in S1 File.
The sequences of the most frequent haplotypes were compared to sequences available in the GenBank database. These sequences shared identity with several sequences identified in humans, animals and even environmental samples. The only exception was the HP01-bg haplotype, which contained 13 identical sequences (100% from the DC group) that did not exactly match any sequence previously deposited in GenBank and was thus considered a novel sequence (Table S14 in S1 File).
Genetic diversity.
Haplotype diversity was evaluated in the four groups. The DC group exhibited considerable haplotype diversity for the gdh (0.91±0.05) and tpi (0.85±0.06) genes. The bg gene revealed the lowest rate (0.34±0.14). The HC group presented rates of 0.71±0.07, 0.85±0.03 and 0.92±0.05 for the gdh, tpi and bg genes, respectively. The ENV and VET groups also presented high levels of haplotype diversity, but this result might be an artifact due to the small sample sizes of these groups.
Multilocus genotyping.
Only 13 samples from the 37 isolates sequenced at the three loci showed no double peaks and presented complete agreement across all genes. The subtypes that were sequenced at all three loci and shared the same assemblage were grouped into 7 MLGs for assemblage A (labeled as BRA1-BRA7), and 3 MLGs for assemblage B isolates (labeled as BRAB1-BRAB3). The MLGs defined for assemblage A (Table 5) corresponded to assemblage AII, and the MLGs defined for assemblage B (Table 6) all corresponded to assemblage BIV. All of these MLGs were obtained from HC and DC isolates. All of the MLGs defined for assemblages A and B and their mutations for each gene are displayed in Tables S15 and S16 in S1 File.
The trees obtained from the concatenated tpi, bg and gdh sequences presented similar results in comparison with the phylogenetic analyses of individual genes and indicated that no recombination events had occurred among these isolates. The topologies observed within different Giardia assemblages confirmed the monophyly of assemblages A and B (Fig. 3). Multilocus genotyping was also performed for samples in which sequences from two genes were obtained. This analysis allowed us to characterize three isolates from assemblage A (Table S17 in S1 File) in the proposed MLGs and to identify four new MLGs from assemblage B (Table S18 in S1 File), including one isolate from an environmental sample. The genetic differences among all genetic assemblages are displayed in Tables S19, S20 and S21 in S1 File.
Phylogenetic relationships of G. duodenalis with Bayesian posterior probabilities using a Markov chain Monte Carlo sampling technique. Markov chains were run for 6,000,000 iterations and the trees were sampled every 500 iterations. The GTR model was used with gamma correction. Sequences from G. muris were employed used as an outgroup.
Discussion
G. duodenalis contamination in the metropolitan region of Campinas
Protozoan contamination has regularly been documented at important environmental sites in the metropolitan region of Campinas. Previous studies have demonstrated the occurrence of Giardia spp. and Cryptosporidium spp. in superficial raw water from the Atibaia River [19], waste water from an effluent treatment plant [57], activated sludge samples from a sewage treatment plant [22], and the intake pipe of a water treatment plant in Campinas [58]. A recent study recognized the metropolitan region of Campinas as presenting the highest risk of Giardia infection among four densely populated regions of Sao Paulo State, Brazil [21]. Likewise, many clinical studies in the same region have reported Giardia infections among children at day-care centers [59] as well as the users of public health services [60], [61], and schools [62] and inhabitants of farms [63]. The great majority of studies only obtained data on the species level based on the morphology of the parasites. Moreover, there is a remarkable lack of studies that correlate environmental contamination with clinical infections in the same region.
Our results demonstrate that there is persistent contamination at important environmental sites in the city of Campinas, such as in streams in the downtown area and rivers (Anhumas River and Atibaia River). This contamination is relevant for people who live in this region. The Atibaia River is impacted by discharge from domestic and industrial sewage and contamination from agricultural sources [58]. The contaminated effluent from the Samambaia Waste Water Treatment Plant represents another source of pathogenic organisms for this river. This effluent is discharged into the Pinheiros Stream, which flows to the Atibaia River. This contamination is important to public health, as several cities use the water of this river for drinking water. The clinical samples obtained in this study (day-care and hospital samples) are discharged into the Anhumas River, which is another tributary of the Atibaia River. These sources of contamination certainly contribute to the high risk of Giardia infection assigned to this metropolitan region [21].
Prevalence of each genetic assemblage in environmental and clinical isolates
Assemblages A and B were the most common in the human clinical samples. These results were expected because people are predominantly infected with these Giardia assemblages. In contrast, assemblage C was identified in five human hospital samples. Although this assemblage has been previously reported in human samples [34], [64], this result can be considered unusual because assemblage C is generally identified in dogs, cats, coyotes and wolves [34]. The detected assemblage C subtypes did not match the other subtypes available in the databases, with the exception of the HC13 isolate, for which an identical sequence was identified in a dog from Sweden [46].
The isolates sequenced from non-human clinical samples belonged mainly to host-adapted assemblages. However, the isolates from this group could be sequenced using the generic tpi marker, revealing the presence of assemblages with zoonotic potential in 83% (5/6) of the veterinary samples, leading us to conclude that these assemblages may present a risk to public health. Although dogs, cats and livestock are usually infected with host-adapted G. duodenalis assemblages [12], [18], these animals can act as potential reservoirs for zoonotic transmission, which might serve as another source of infection for people living in this region.
In the present study, most of the environmental samples were characterized as belonging to assemblages A and B, including the waste water sample. Similar results have been obtained at other waste water treatment plants in other countries, such as China [65], the United States [66] and France [67]. These results suggest that anthroponotic transmission is important in these environmental sources, although the zoonotic potential of these samples must be considered. Assemblage D was also observed in the Anhumas River, and assemblage C was identified at the water abstraction site of the city. These findings suggest that there are other sources of contamination at these environmental sites. The identification of Giardia cysts in water supplies may contribute substantially to the protection of public health in the investigated areas [68].
Mixed assemblages and heterogeneous sequence profiles
In the present work, mixed assemblage isolates were observed in the human hospital samples (12%) and a remarkably high rate of such assemblages (43%) was noted in children from the day-care center. It is also noteworthy that in all of the mixed assemblages obtained from the day-care center, all bg sequences were identified as assemblage AII, whereas the other markers indicated assemblage BIV or BIII. This difference may be due to mixed infections showing preferential amplification of single genotypes over others at a specific locus [69]. Mixed assemblage isolates have been frequently reported in studies that make use of more than a single molecular marker [34], [69]–[71]. The finding that most of the mixed assemblages identified in this study were from assemblages A and B can be explained by the overrepresentation of human samples, which were predominantly assigned to these assemblages.
Despite the small number of veterinary samples examined, mixed assemblages were found in all of the veterinary samples sequenced with at least two markers. This result suggests that there must be high endemicity of Giardia in these animals and that they likely continue to re-infect themselves due to contact with the feces of other infected animals. Further studies are required to determine the prevalence rates of Giardia duodenalis and the most common G. duodenalis assemblages in both household and kennel animals in this metropolitan region.
Overlapping nucleotides were identified at a high frequency with similar rates among all three markers. Similar results were found in lambs from Spain, in which the frequency of mixed templates reached 44% for the tpi gene [72]. A study of human and animal samples conducted in Italy, Africa and Croatia detected rates of overlapping nucleotides of 36%, 50% and 75% in the sequences of the bg, tpi and gdh genes, respectively [73]. Several studies have reported a high percentage of double peaks in the chromatograms in various animal species, including humans [46], [74], [75].
Three cloned samples in which more than one haplotype had been previously detected, presented different subtypes in their clones. Two samples (VET02 and VET04) exhibited different genetic assemblages in their clones. Sample HC43 displayed different subtypes, all from the same genetic assemblage. After sequencing of the clones, these sequences were found to present more polymorphisms than had been detected in the double peaks. These results suggest that most sequences showing double peaks previously identified at specific sites most likely represent mixed assemblages, rather than allelic sequence heterozygosity.
The uncertainty regarding the origin of the mixed assemblages and/or mixed templates is due to the difficulty associated with determining whether the mixed profile arose through genetic exchange, allelic polymorphisms, mixed/concurrent infections or a combination of all of these events. Higher rates of mixed assemblages were observed in the DC and VET groups. Because there is close contact among children at day-care centers and among infected dogs and cats in the streets, these results likely represent mixed infections.
Haplotype diversity and zoonotic potential
The main importance of the haplotypes analysis lies in the observation that the number of isolates found in each genetic assemblage is not necessarily followed by a proportional number of haplotypes depending on the genetic diversity of each group of samples or the molecular markers used. In the present study, the haplotype analysis of the tpi and bg genes indicated that there was higher variability in samples of assemblage B, which is consistent with previous studies investigating sequence polymorphisms in assemblage B compared with assemblage A [76], [77].
A considerable proportion of different haplotypes was found at all of the sources from which positive samples were obtained. The haplotype variability observed in the hospital samples (48 haplotypes) can be explained by both the different origins of people who attend these hospitals and the overall high genetic variability of the human samples classified into assemblages A and B. Patients at these hospitals come not only from the metropolitan region but also from the state of São Paulo. The pronounced variability in the HC group most likely reflects the haplotypes of the entire state. Further molecular surveillance studies are required to verify the extent of hospital sewage contamination in the water sources for the metropolitan region, as this sewage may be introducing new Giardia haplotypes into environmental sites in this region.
The haplotype variability detected in the samples from the day-care center (31 haplotypes) may be a consequence of the multiple sources of infection observed at this location combined with the fact that children attending this center live in conditions of socioeconomic vulnerability and experience unsanitary crowding conditions. Several potential sources of zoonosis (including dogs and cats) have been observed in the vicinity of the day-care center, and these species may have served as reservoirs for zoonotic transmission. The presence of some haplotypes with high frequencies suggests that these haplotypes may have been introduced into the day-care center population through person-to-person transmission.
Analyses of the VET and ENV groups revealed a regular distribution of haplotypes with similar frequencies. The VET group consists of different animals with infections that likely originated from different sources. Our results for the ENV group suggest that many different haplotypes are present at different environmental sites. These sites are vulnerable to many sources of contamination, such as hospital and domestic sewage, animal feces and agricultural and industrial sewage. All of these sources may contribute to the observed haplotype variability, and further studies are necessary to monitor the most common haplotypes of Giardia at each environmental site.
Although a considerable number of haplotypes were identified, most of the samples originated from only a few haplotypes. Thus, only a small proportion of the isolates were responsible for the large number of haplotypes detected. The observation of a particular haplotype in different groups suggests potential relationships among these groups, which is consistent with the previously demonstrated interconnections of the collection sites. Determination of whether one group is contaminating all of the others or if they are all vulnerable to the same sources of contamination, will require further molecular surveillance studies.
Characterization of the genetic profile of G. duodenalis in the metropolitan region via multilocus genotyping
In the present study, 10 different MLGs were identified in the isolates from the hospitals and day-care center. With the exception of the BRA1 MLG, all other MLGs corresponded to new profiles. The identified MLGs may contribute to future studies designed to establish a genetic profile of the most common genotypes found in this region. A secondary analysis, including isolates that lacked one of the three gene sequences, allowed us to identify new profiles, including a new MLG originating from an environmental sample. Characterization of environmental isolates should play an important role in future studies given the absence of information about contributors to environmental contamination [73].
Final remarks
In conclusion, in addition to Giardia contamination at important environmental sites and infections in non-human hosts, our data demonstrate that most of the Giardia isolates obtained in this work can be considered to show zoonotic potential. There was a high degree of genetic diversity in the isolates obtained from both clinical and environmental samples, suggesting that multiple sources of infection are responsible for the detected contamination events. The detection of different assemblages and subtypes (after molecular cloning) in isolates with and without double peaks suggests that most studies likely underestimate the real genetic diversity of this parasite.
Our results, together with those of previous studies reporting environmental contamination and prevalence of Giardia above the limits tolerated by the United States Environmental Protection Agency (USEPA) [19], [21], [22], indicate a notable risk for human infection by Giardia in this region.
Future molecular surveillance studies of G. duodenalis and other waterborne protozoans are essential to elucidate the genetic profile of the distribution of these parasites in this region. Water sources, waste water effluents, water treatment plants, sewage treatment plants, hospitals, day-care centers, and public kennels are some key potential study sites. Analyses of population genetic structure conducted with high-resolution tools are needed to elucidate additional features of the origin and dissemination of the parasite [78]. Complementary approaches are necessary to detect mixed sequence profiles and rare haplotypes to avoid underestimation of the genetic diversity of Giardia, such as a) molecular cloning and sequencing of amplified fragments, b) the use of assemblage-specific primers, and c) high-throughput next-generation sequencing. The multilocus genotype approach presented herein can be considered to represent an initial step in the characterization of Giardia diversity in the metropolitan region of Campinas and can help to track sources of contamination associated with this waterborne protozoan. Furthermore, to prevent the spread of waterborne diseases, new public policies should consider not only the city but also the complex relationships among different cities that share the same metropolitan region.
Supporting Information
S1 Fig.
Phylogenetic analysis of the VET group. Phylogenetic analysis of the Giardia triose phosphate isomerase (tpi) gene from the VET group. The alignment was generated with ClustalX and analyzed using maximum likelihood with the Tamura-Nei 93 model with gamma correction (MEGA v5.05). Trees derived using neighbor-joining produced a similar topology. Bootstrap values>50% (10,000 replicates) are shown beside each node. Accession numbers for tpi reference sequences are shown beside the corresponding assemblage.
https://doi.org/10.1371/journal.pone.0115489.s001
(TIF)
S1 File.
Table S1, Accession numbers for gene sequences obtained from NCBI. All reference sequences used in this study from the three genes are listed in this table. Table S2, Molecular characterization and accession numbers of isolates obtained from hospitals based on sequencing data from the gdh, bg, and tpi genes. The genetic assemblage of each sequenced isolate from the HC group and its accession numbers are displayed in this table. Table S3, Molecular characterization and accession numbers of isolates obtained from the day-care center based on sequencing data from the gdh, bg, and tpi genes. The genetic assemblage of each sequenced isolate in the DC group and its accession numbers are displayed in this table. Table S4, Molecular characterization and accession numbers of isolates obtained from veterinary samples based on sequencing data from the gdh, bg, and tpi genes. The genetic assemblage of each sequenced isolate from the VET group and its accession numbers are displayed in this table. Table S5, Molecular characterization and accession numbers of isolates obtained from environmental samples based on sequencing data from the gdh, bg, and tpi genes. The genetic assemblage of each sequenced isolate from the ENV group and its accession numbers are displayed in this table. Table S6, Distribution of each source as a percentage within each genetic assemblage. Each assemblage was examined to identify the source that contributed the most isolates. The table presents the proportion of each source of contamination (HC, DC, VET and ENV) within each genetic assemblage. Table S7, Samples with double peaks within each loci. All samples that presented double peaks are listed based on sequencing data from the gdh, bg, and tpi genes. Table S8, Correspondence between subtypes and sequences of the bg gene in the isolates. Correspondence between subtypes of the bg gene and the associated isolates. Each subtype (S) presents at least one isolate. The main subtypes revealed many isolates in one subtype. Table S9, Correspondence between the subtypes and sequences of the gdh gene in the isolates. Correspondence between subtypes of the gdh gene and the associated isolates. Each subtype (S) presents at least one isolate. The main subtypes revealed many isolates in one subtype. Table S10, Correspondence between subtypes and sequences of the tpi gene in the isolates. Correspondence between subtypes of the tpi gene and the associated isolates. Each subtype (S) presents at least one isolate. The main subtypes revealed many isolates in one subtype. Table S11, Correspondence between the haplotypes and sequences of the gdh gene in the isolates. Correspondence between haplotypes of the gdh gene and the associated isolates. The main haplotypes presents many isolates with each haplotype. Some haplotypes showed a mixture among the different groups. Table S12, Correspondence between the haplotypes and sequences of the tpi gene in the isolates. Correspondence between the haplotypes of the tpi gene and the associated isolates. The main haplotypes present many isolates with each haplotype. Some haplotypes showed a mixture among the different groups. Table S13, Correspondence between the haplotypes and sequences of the bg gene in the isolates. Correspondence between the haplotypes of the bg gene and the associated isolates. The main haplotypes present many isolates with each haplotype. Some haplotypes revealed a mixture among the different groups. Table S14, Comparison of each haplotype identified with sequences available in the GenBank database. This table shows the main haplotypes obtained in this study. These haplotypes were compared with the GenBank database. Many identical sequences were identified. The haplotype HP01-bg was not identified in the database and corresponds to a new sequence. Table S15, Polymorphic sites in the bg and tpi sequences among G. duodenalis assemblage A isolates. This table presents all mutations that contributed to differences in each multilocus genotypes from assemblage A isolates. The gdh gene did not present any differences among the multilocus genotypes identified. Table S16, Polymorphic sites in the bg, tpi, and gdh sequences among G. duodenalis assemblage A isolates. This table presents all mutations that contributed to differences in each multilocus genotype from assemblage B isolates. Table S17, Multilocus genotypes (MLGs) in G. duodenalis assemblage A isolates. Multilocus genotypes obtained from assemblage A isolates in which at least two sequences were obtained. Table S18, Multilocus genotypes (MLGs) in G. duodenalis assemblages B isolates. Multilocus genotypes obtained from assemblage B isolates in which at least two sequences were obtained. Table S19, Genetic differences in the genetic assemblages at the gdh locus. All SNPs in the different genetic assemblages are displayed within gdh loci. Table S20, Genetic differences in the genetic assemblages at the bg locus. All SNPs in the different genetic assemblages are displayed within bg loci. Table S21, Genetic differences in the genetic assemblages at the tpi locus. All SNPs in the different genetic assemblages are displayed within tpi loci.
https://doi.org/10.1371/journal.pone.0115489.s002
(DOCX)
Acknowledgments
We thank Dr. Carlos Emílio Levy, head of the Clinical Pathology Division of the University of Campinas Clinic Hospital; Edson Gonçalves (Animaltec); and Drs. Diego A. G. Leal, Juliane A. G. Araújo, Luciana U. Santos and Jancarlo Ferreira Gomes for generously providing samples of Giardia cysts. We thank the Sociedade de Abastecimento de Água e Saneamento S.A. (SANASA) and Dr. Romeu Cantúsio Neto for providing a sample from the water abstraction site.
Author Contributions
Conceived and designed the experiments: MD APS. Performed the experiments: MD. Analyzed the data: MD AGA MIZ RMBF. Contributed reagents/materials/analysis tools: MD APS. Wrote the paper: MD AGA MIZ RMBF APS.
References
- 1. Adam RD (2001) Biology of Giardia lamblia. Clin Microbiol Rev 14:447–475.
- 2. Cavalier-Smith T (2004) Only six kingdoms of life. Proc Biol Sci 271:1251–1262.
- 3. Adl SM, Simpson AGB, Farmer MA, Andersen RA, Anderson OR, et al. (2005) The new higher level classification of eukaryotes with emphasis on the taxonomy of protists. J Eukaryot Microbiol 52:399–451.
- 4. Yoder JS, Harral C, Beach MJ (2012) Giardiasis Surveillance — United States, 2009–2010. MMWR Surveill Summ 59:15–25.
- 5. Ankarklev J, Hestvik E, Lebbad M, Lindh J, Kaddu-Mulindwa DH, et al. (2012) Common coinfections of Giardia intestinalis and Helicobacter pylori in non-symptomatic Ugandan children. PLoS Negl Trop Dis 6:e1780.
- 6. Savioli L, Smith H, Thompson A (2006) Giardia and Cryptosporidium join the “Neglected Diseases Initiative”. Trends Parasitol 22:203–208.
- 7.
CDC (1994) Addressing Emerging Infectious Disease Threats: A Prevention Strategy for the United States. Atlanta.
- 8.
WHO (2002) The World Health Report 2002 – reducing risks, promoting healthy life. World Health Organization. Geneva, Switzerland.
- 9.
WHO (2011) The World Health Statistics 2011. Geneva, World Health Organization. Geneva, Switzerland.
- 10. Thompson RCA, Palmer CS, O'Handley R (2008) The public health and clinical significance of Giardia and Cryptosporidium in domestic animals. Vet J 177:18–25.
- 11. Paz e Silva FM, E Silva P, Monobe MM, Lopes RS, Araujo JP (2012) Molecular characterization of Giardia duodenalis in dogs from Brazil. Parasitol Res 110:325–334.
- 12. Bowman DD, Lucio-Forster A (2010) Cryptosporidiosis and giardiasis in dogs and cats: veterinary and public health importance. Exp Parasitol 124:121–127.
- 13. Palmer CS, Traub RJ, Robertson ID, Devlin G, Rees R, et al. (2008) Determining the zoonotic significance of Giardia and Cryptosporidium in Australian dogs and cats. Vet Parasitol 154:142–147.
- 14.
Paoletti B, Otranto D, Weigl S, Giangaspero A, Cesare A Di, et al. (2010) Prevalence and genetic characterization of Giardia and Cryptosporidium in cats from Italy. Res Vet Sci: 10–12.
- 15. Geurden T, Vanderstichel R, Pohle H, Ehsan A, von Samson-Himmelstjerna G, et al. (2012) A multicentre prevalence study in Europe on Giardia duodenalis in calves, with molecular identification and risk factor analysis. Vet Parasitol 190:383–390.
- 16. Geurden T, Levecke B, Pohle H, De Wilde N, Vercruysse J, et al. (2010) A Bayesian evaluation of two dip-stick assays for the on-site diagnosis of infection in calves suspected of clinical giardiasis. Vet Parasitol 172:337–340.
- 17.
O'Handley RM (2002) Giardia in farm animals. In Giardia: The cosmopolitan parasite. Olson EB, Olson M, Wallis PM, editors London, UK.: CAB International.
- 18. Zhang W, Zhang X, Wang R, Liu A, Shen Y, et al. (2012) Genetic characterizations of Giardia duodenalis in sheep and goats in Heilongjiang Province, China and possibility of zoonotic transmission. PLoS Negl Trop Dis 6:e1826.
- 19. Franco RMB, Rocha-Eberhardt R, Neto RC (2001) Occurrence of Cryptosporidium oocysts and Giardia cysts in raw water from the Atibaia river, Campinas, Brazil. Rev Inst Med Trop Sao Paulo 43:109–111.
- 20. Haramoto E, Kitajima M, Kishida N, Katayama H, Asami M, et al. (2012) Occurrence of viruses and protozoa in drinking water sources of Japan and their relationship to indicator microorganisms. Food Environ Virol 4:93–101.
- 21. Sato MIZ, Galvani AT, Padula JA, Nardocci AC, Lauretto MDS, et al. (2013) Assessing the infection risk of Giardia and Cryptosporidium in public drinking water delivered by surface water systems in Sao Paulo State, Brazil. Sci Total Environ 442:389–396.
- 22. Santos LU, Bonatti TR, Neto RC, Franco RMB (2004) Occurrence of Giardia cysts and Cryptosporidium oocysts in activated sludge samples in Campinas, SP, Brazil. Rev Inst Med Trop Sao Paulo 46:309–313.
- 23. Monis PT, Andrews RH, Mayrhofer G, Mackrill J, Kulda J, et al. (1998) Novel lineages of Giardia intestinalis identified by genetic analysis of organisms isolated from dogs in Australia. Parasitology 116 Pt 1:7–19.
- 24. Andrews RH, Adams M, Boreham PFL, Melonis BP (1989) Giardia intestinalis: Electrophoretic Evidence for A Species Complex. Int J Parasitol 19:183–190.
- 25. Homan WL, Enckevort FHJ, Limper L, Eys GJJM, Schoone GJ, et al. (1992) Comparison of Giardia isolates from different laboratories by isoenzyme analysis and recombinant DNA probes. Parasitol Res 78:316–323.
- 26. Caccio SM, Sprong H (2010) Giardia duodenalis: genetic recombination and its implications for taxonomy and molecular epidemiology. Exp Parasitol 124:107–112.
- 27. Monis PT, Mayrhofer G, Andrews RH, Homan WL, Limper L, et al. (1996) Molecular genetic analysis of Giardia intestinalis isolates at the glutamate dehydrogenase locus. Parasitology 112 Pt 1:1–12.
- 28. Hopkins RM, Meloni BP, Groth DM, Wetherall JD, A J, et al. (1997) Ribosomal RNA Sequencing Reveals Differences between the Genotypes of Giardia Isolates Recovered from Humans and Dogs Living in the Same Locality. J Parasitol 83:44–51.
- 29. Nash TE, Mowatt MR (1992) Identification and characterization of a Giardia lamblia group-specific gene. Exp Parasitol 75:369–378.
- 30. Zhang W, Shen Y, Wang R, Liu A, Ling H, et al. (2012) Cryptosporidium cuniculus and Giardia duodenalis in rabbits: genetic diversity and possible zoonotic transmission. PLoS One 7:e31262.
- 31. Monis PT (1999) The importance of systematics in parasitological research. Int J Parasitol 29:381–388.
- 32. Caccio SM, Ryan U (2008) Molecular epidemiology of giardiasis. Mol Biochem Parasitol 160:75–80.
- 33. Thompson RCA, Hopkins RM, Homan WL (2000) Nomenclature and Genetic Groupings of Giardia Infecting Mammals Early taxonomy. Parasitology 16:0–3.
- 34. Sprong H, Der JWB Van, Caccio SM (2009) Identification of Zoonotic Genotypes of Giardia duodenalis. PLoS Negl Trop Dis 3:1–12.
- 35. Lasek-Nesselquist E, Welch DM, Sogin ML (2010) The identification of a new Giardia duodenalis assemblage in marine vertebrates and a preliminary analysis of G. duodenalis population biology in marine systems. Int J Parasitol 40:1063–1074.
- 36. Faust EC, D'Antoni JS, Odom V, Miller MJ, Peres C, et al. (1938) A Critical Study of Clinical Laboratory Technics for the Diagnosis of Protozoan Cysts and Helminth Eggs in Feces: I. Preliminary Communication. Am J Trop Med Hyg s1-18:169–183.
- 37. Smith AL, Smith HV (2009) A comparison of fluorescein diacetate and propidium iodide staining and in vitro excystation for determining Giardia intestinalis cyst viability. Parasitology 99:329.
- 38. Shepherd KM, Wyn-Jones a P (1996) An evaluation of methods for the simultaneous detection of Cryptosporidium oocysts and Giardia cysts from water. Appl Environ Microbiol 62:1317–1322.
- 39. Neto RC, Santos LU, Franco RMB (2006) Evaluation of activated sludge treatment and the efficiency of the disinfection of Giardia species cysts and Cryptosporidium oocysts by UV at a sludge treatment plant in Campinas, south-east Brazil. Water Sci Technol 54:89.
- 40. Mahbubani MH, Bej a K, Perlin MH, Schaefer FW, Jakubowski W, et al. (1992) Differentiation of Giardia duodenalis from other Giardia spp. by using polymerase chain reaction and gene probes. J Clin Microbiol 30:74–78.
- 41.
Wielinga CM, Ryan U, Andrew Thompson RC, Monis P (2011) Multi-locus analysis of Giardia duodenalis intra-Assemblage B substitution patterns in cloned culture isolates suggests sub-Assemblage B analyses will require multi-locus genotyping with conserved and variable genes. Int J Parasitol: 1–9.
- 42. Sulaiman IM, Fayer R, Bern C, Gilman RH, Trout JM, et al. (2003) Triosephosphate isomerase gene characterization and potential zoonotic transmission of Giardia duodenalis. Emerg Infect Dis 9:1444–1452.
- 43. Abe N, Kimata I, Iseki M (2003) Identification of genotypes of Giardia intestinalis isolates from dogs in Japan by direct sequencing of the PCR amplified glutamate dehydrogenase gene. J Vet Med Sci 65:29–33.
- 44. Caccio SM, De Giacomo M, Pozio E (2002) Sequence analysis of the beta-giardin gene and development of a polymerase chain reaction-restriction fragment length polymorphism assay to genotype Giardia duodenalis cysts from human faecal samples. Int J Parasitol 32:1023–1030.
- 45. Kreader Ca (1996) Relief of amplification inhibition in PCR with bovine serum albumin or T4 gene 32 protein. Appl Environ Microbiol 62:1102–1106.
- 46. Lebbad M, Mattsson JG, Christensson B, Ljungström B, Backhans A, et al. (2010) From mouse to moose: multilocus genotyping of Giardia isolates from various animal species. Vet Parasitol 168:231–239.
- 47. Larkin Ma, Blackshields G, Brown NP, Chenna R, McGettigan Pa, et al. (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23:2947–2948.
- 48. Tamura K, Peterson D, Peterson N, Stecher G, Nei M, et al. (2011) MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods. Mol Biol Evol 28:2731–2739.
- 49. Posada D (2008) jModelTest: phylogenetic model averaging. Mol Biol Evol 25:1253–1256.
- 50. Tamura K, Nei M (1993) Estimation of the number of nucleotide substitutions in the control region of mitochondrial DNA in humans and chimpanzees. Mol Biol Evol 10:512–526.
- 51. Huelsenbeck JP, Ronquist F (2001) MRBAYES: Bayesian inference of phylogenetic trees. Bioinformatics 17:754–755.
- 52. Page RD (1996) TreeView: an application to display phylogenetic trees on personal computers. Comput Appl Biosci 12:357–358.
- 53. Maddison DR, Swofford DL, Maddison WP (1997) NEXUS: an extensible file format for systematic information. Syst Biol 46:590–621.
- 54. Clement M, Posada D, Crandall Ka (2000) TCS: a computer program to estimate gene genealogies. Mol Ecol 9:1657–1659.
- 55. Rozas J, Sanchez-DelBarrio JC, Messeguer X, Rozas R (2003) DnaSP, DNA polymorphism analyses by the coalescent and other methods. Bioinformatics 19:2496–2497.
- 56. Excoffier L, Lischer HEL (2010) Arlequin suite ver 3.5: a new series of programs to perform population genetics analyses under Linux and Windows. Mol Ecol Resour 10:564–567.
- 57. Santos LU, Alves DP, Guaraldo AMA, Neto RC, Durigan M, et al. (2013) Infectivity of Giardia duodenalis cysts from UV light-disinfected wastewater effluent using a nude BALB/c mouse model. ISRN Parasitol 2013:1–7.
- 58. Neto RC, dos Santos LU, Sato MIZ, Franco RMB (2010) Cryptosporidium spp. and Giardia spp. in surface water supply of Campinas, southeast Brazil. Water Sci Technol 62:217–222.
- 59. Franco RMB, Cordeiro N da S (1996) Giardiose e Criptosporidiose em Creches no Município de Campinas, SP. Rev Soc Bras Med Trop 29:585–591.
- 60. Cardoso FM, Morcillo AM, Zeferino AMB, Antonio MÂR de GM (2005) Enteroparasites in users of the public health services in the city of Campinas, São Paulo, Brazil. Rev Ciências Médicas 14:337–343.
- 61. Gioia I (1992) Prevalência de parasitoses intestinais entre os usuários do centro de saúde do distrito de Sousas, Campinas, SP. Rev Soc Bras Med Trop 25:177–182.
- 62. Moura ECde, Bragazza LM, Coelho MFL, Au SMF (1997) Prevalence of intestinal parasitosis in schoolchildren. J Pediatr (Rio J) 73:406–410.
- 63. Kobayashi J, Hasegawa H, Forli a a, Nishimura NF, Yamanaka A, et al. (1995) Prevalence of intestinal parasitic infection in five farms in Holambra, São Paulo, Brazil. Rev Inst Med Trop Sao Paulo 37:13–18.
- 64. Soliman RH, Fuentes I, Rubio JM (2011) Identification of a novel Assemblage B subgenotype and a zoonotic Assemblage C in human isolates of Giardia intestinalis in Egypt. Parasitol Int 60:507–511.
- 65. Li N, Xiao L, Wang L, Zhao S, Zhao X, et al. (2012) Molecular surveillance of Cryptosporidium spp., Giardia duodenalis, and Enterocytozoon bieneusi by genotyping and subtyping parasites in wastewater. PLoS Negl Trop Dis 6:e1809.
- 66.
Sulaiman IM, Jiang J, Singh A, Xiao L (2004) Distribution of Giardia duodenalis Genotypes and Subgenotypes in Raw Urban Wastewater in Milwaukee, Wisconsin Distribution of Giardia duodenalis Genotypes and Subgenotypes in Raw Urban Wastewater in Milwaukee, Wisconsin.
- 67. Bertrand I, Schwartzbrod J (2007) Detection and genotyping of Giardia duodenalis in wastewater: relation between assemblages and faecal contamination origin. Water Res 41:3675–3682.
- 68. Plutzer J, Karanis P, Domokos K, Törökné A, Márialigeti K (2008) Detection and characterisation of Giardia and Cryptosporidium in Hungarian raw, surface and sewage water samples by IFT, PCR and sequence analysis of the SSUrRNA and GDH genes. Int J Hyg Environ Health 211:524–533.
- 69. Read C, Monis PT, Thompson RCA (2004) Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol 4:125–130.
- 70. Levecke B, Geldhof P, Claerebout E, Dorny P, Vercammen F, et al. (2009) Molecular characterisation of Giardia duodenalis in captive non-human primates reveals mixed assemblage A and B infections and novel polymorphisms. Int J Parasitol 39:1595–1601.
- 71. Al-Mohammed HI (2011) Genotypes of Giardia intestinalis clinical isolates of gastrointestinal symptomatic and asymptomatic Saudi children. Parasitol Res 108:1375–1381.
- 72. Gómez-Muñoz MT, Cámara-Badenes C, Martínez-Herrero MDC, Dea-Ayuela MA, Pérez-Gracia MT, et al. (2012) Multilocus genotyping of Giardia duodenalis in lambs from Spain reveals a high heterogeneity. Res Vet Sci 93:836–842.
- 73. Caccio SM, Beck R, Lalle M, Marinculic a, Pozio E (2008) Multilocus genotyping of Giardia duodenalis reveals striking differences between assemblages A and B. Int J Parasitol 38:1523–1531.
- 74. Geurden T, Thomas P, Casaert S, Vercruysse J, Claerebout E (2008) Prevalence and molecular characterisation of Cryptosporidium and Giardia in lambs and goat kids in Belgium. Vet Parasitol 155:142–145.
- 75. Lebbad M, Petersson I, Karlsson L, Botero-Kleiven S, Andersson JO, et al. (2011) Multilocus genotyping of human Giardia isolates suggests limited zoonotic transmission and association between assemblage B and flatulence in children. PLoS Negl Trop Dis 5:e1262.
- 76. Franzén O, Jerlström-Hultqvist J, Castro E, Sherwood E, Ankarklev J, et al. (2009) Draft genome sequencing of Giardia intestinalis assemblage B isolate GS: is human giardiasis caused by two different species? PLoS Pathog 5:e1000560.
- 77. Lebbad M, Ankarklev J, Tellez A, Leiva B, Andersson JO, et al. (2008) Dominance of Giardia assemblage B in León, Nicaragua. Acta Trop 106:44–53.
- 78. Wang R, Zhang L, Axén C, Bjorkman C, Jian F, et al. (2014) Cryptosporidium parvum IId family: clonal population and dispersal from Western Asia to other geographical regions. Sci Rep 4:4208.