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Molecular epidemiologic tools for waterborne pathogens Cryptosporidium spp. and Giardia duodenalis

Lihua Xiao, Yaoyu Feng

Food and Waterborne Parasitology · 2017 · Review · CC BY

doi:10.1016/j.fawpar.2017.09.002 (opens in a new tab)

Abstract

Molecular diagnostic tools have played an important role in improving our understanding of the transmission of Cryptosporidium spp. and Giardia duodenalis, which are two of the most important waterborne parasites in industrialized nations. Genotyping tools are frequently used in the identification of host-adapted Cryptosporidium species and G. duodenalis assemblages, allowing the assessment of infection sources in humans and public health potential of parasites found in animals and the environment. In contrast, subtyping tools are more often used in case linkages, advanced tracking of infections sources, and assessment of disease burdens attributable to anthroponotic and zoonotic transmission. More recently, multilocus typing tools have been developed for population genetic characterizations of transmission dynamics and delineation of mechanisms for the emergence of virulent subtypes. With the recent development in next generation sequencing techniques, whole genome sequencing and comparative genomic analysis are increasingly used in characterizing Cryptosporidium spp. and G. duodenalis. The use of these tools in epidemiologic studies has identified significant differences in the transmission of Cryptosporidium spp. in humans between developing countries and industrialized nations, especially the role of zoonotic transmission in human infection. Geographic differences are also present in the distribution of G. duodenalis assemblages A and B in humans. In contrast, there is little evidence for widespread zoonotic transmission of giardiasis in both developing and industrialized countries. Differences in virulence have been identified among Cryptosporidium species and subtypes, and possibly between G. duodenalis assemblages A and B, and genetic recombination has been identified as one mechanism for the emergence of virulent C. hominis subtypes. These recent advances are providing insight into the epidemiology of waterborne protozoan parasites in both developing and developed countries.

Full text

From PubMed Central, U.S. National Library of Medicine, as current on Oct 20, 2026. Paragraph numbers are ours, so an answer can point at one.

Introduction

Protozoan parasites are important causes of diarrhea and other enteric diseases in humans. They include mostly Cryptosporidium spp., Giardia duodenalis, Cyclospora cayetanensis, Cystoisospora belli, Entamoeba histolytica, and Toxoplasma gondii, although human infections with the latter are usually associated with non-gastrointestinal symptoms. Other protozoa in humans are not considered major pathogens, such as other Entamoeba spp., Blastocystis hominis, and Balantidium coli. As these pathogens use the fecal-oral route to maintain the lifecycle, foodborne and waterborne transmission are important in disease epidemiology. Among them, Cryptosporidium spp. and G. duodenalis are best known for their potential to cause large waterborne outbreaks of illness, thus are major targets in recent research on waterborne protozoan pathogens in humans. Earlier waterborne outbreaks of diseases by these pathogens were mostly associated with drinking water, but more recently, recreational water-associated outbreaks of cryptosporidiosis and giardiasis are increasingly reported, and for cryptosporidiosis, they are now responsible for most waterborne outbreaks in the United States.

Molecular diagnostic tools have long been used in studies of the transmission of Cryptosporidium spp. and G. duodenalis. This is largely due to the existence of a large number of morphologically identical species or genotypes within both group of protozoa, which require the use of molecular diagnostic tools for differentiation. Thus far, over 30 Cryptosporidium species and many genotypes of unknown species status have been described. Similarly, there are at least eight genotypes (assemblages) of G. duodenalis that are likely cryptic species. Because of the existence of host specificity among Cryptosporidium species and G. duodenalis genotypes, characterizations of pathogens at the species or genotype level are helpful to the assessment of infection sources in humans and public health potential of parasites in animals and the environment. Advanced characterization of human-pathogenic species or genotypes can be used in case linkage, tracking of virulent subtypes, and assessment of disease burdens attributable to different transmission routes. This can be achieved through conventional subtyping based on sequence analysis of individual polymorphic genes, multilocus typing, and comparative genomic analysis. In this report, we will review various molecular diagnostic tools used in the characterization of human-pathogenic Cryptosporidium spp. and G. duodenalis.

Genotyping tools for Cryptosporidium spp.

Accurate identification of Cryptosporidium species requires the use of genotyping tools. Currently, most Cryptosporidium genotyping tools use PCR targeting the small subunit (SSU) rRNA gene, largely because of the existence of conserved Cryptosporidium-specific sequence for designing primers that allow broad specific detection of all Cryptosporidium spp. and semi-conserved and hypervariable regions that can be used for the differentiation of various species and genotypes by restriction fragment length polymorphism (RFLP), melting curve, or DNA sequence analyses. PCR tools targeting other genes were used in early Cryptosporidium research, but they are now infrequently used in Cryptosporidium genotyping because of their narrow detection range; they can only be used for genotyping Cryptosporidium species that are closely related to C. parvum or C. hominis because of the nature of sequences used in primer design.

Among the SSU rRNA-based Cryptosporidium genotyping tools, the PCR-RFLP tool using nested PCR amplification of a ~ 830-bp fragment and restriction analysis of the secondary PCR products using enzymes SspI and VspI is the most commonly used one. It can be used in genotyping Cryptosporidium spp. from both humans and animals. More recently, the use of VspI has been replaced with MboII in RFLP analysis of PCR products from Cryptosporidium spp. in ruminants. The technique requires stringent PCR conditions, but when optimized, can be used effectively in detecting single oocysts in water samples. Alternatively, a PCR assay that targeting a smaller fragment of the SSU rRNA gene can be used. The technique is more adaptable to other PCR buffers, although is slightly less specific for Cryptosporidium (mostly presenting a minor problem to the analysis of water samples) and requires DNA sequence analysis in Cryptosporidium genotype determination.

In recent years, qPCR assays are increasingly used in genotyping human-pathogenic Cryptosporidium spp. Several species-specific genotyping tools based on the SSU rRNA and other genes have been developed for C. hominis, C. parvum, and C. cuniculus. One major issue is the broad range of Cryptosporidium species that can occur in humans, which has limited the wide use of these species-specific qPCR assays. However, some of these tools also include a SSU rRNA-based generic qPCR assay for the detection of all Cryptosporidium species, which can be used in conjunction with C. parvum and C. hominis-specific assays for rapid differentiation of the two dominant Cryptosporidium species in human specimens. Recently, using fluorescence resonance energy transfer probes and melt curve analysis, one SSU rRNA-based PCR assays has been developed for rapid genotyping to five common Cryptosporidium species in human specimens. In addition to the potential of streamlining the detection and identification of Cryptosporidium spp. in human clinical specimens, qPCR-based genotyping assays have been used in quantifying Cryptosporidium oocysts in these specimens, although low levels of oocysts that are typically found in drinking water samples cannot be accurately quantified.

Next generation sequencing techniques are now increasingly used in Cryptosporidium genotyping, especially accurate identifications of mixed Cryptosporidium genotypes. Their usage in sequencing SSU rRNA and actin PCR products from several Cryptosporidium species of humans and animals was effectively demonstrated. Although they are more expensive than traditional Sanger sequencing when analyzing small numbers of specimens, with the use of product indexing, the costs are comparable when specimens are processed in batches. With standardization, these techniques are likely to be used in routine Cryptosporidium genotyping in future.

The use of genotyping tools in the analysis of clinical specimens has led to the identification of over 20 Cryptosporidium species and genotypes in humans (Table 1). With the exception of C. meleagridis, which infects both birds and mammals, all other human-pathogenic Cryptosporidium species and genotypes are pathogens of mammals. It is likely that some other mammalian Cryptosporidium species and genotypes will be found in humans in future. Clearly, these species and genotypes differ significantly in human infectivity, reflected by the differences in their frequency in both sporadic human cases and outbreaks. Molecular epidemiologic studies have also identified differences in the distribution of Cryptosporidium species and genotypes in humans between developing countries and industrialized nations. Thus, C. hominis is the dominant species in humans in developing countries, whereas both C. hominis and C. parvum are common in humans in most industrialized nations. Other species, such as C. meleagridis, C. felis, C. canis, C. ubiquitum, C. cuniculus, C. muris and chipmunk genotype I are mainly animal pathogens, with the exception of C. viatorum, which appears to be exclusively a human pathogen. These species are responsible for variable numbers of human infections depending on socioeconomic development and intensity of agriculture. Among them, C. meleagridis, C. canis, C. viatorum and C. muris are mostly found in children and HIV + persons in developing countries, whereas C. ubiquitum, C. cuniculus, and chipmunk genotype I are mainly found in immunocompetent and immunocompromised persons in industrialized nations. As these Cryptosporidium species and genotypes have different host ranges, the difference in Cryptosporidium species distribution could be a reflection of differences in the importance of various transmission routes in cryptosporidiosis epidemiology. The remaining Cryptosporidium species or genotypes have been reported in only a few patients (Table 1). In addition to differences in human infectivity, Cryptosporidium species differ from each other in virulence, with C. hominis more virulent and causing more clinical presentations in most studies. In a few studies, however, patients infected C. parvum had longer duration of diarrhea or higher occurrence of vomiting than those with C. hominis,.

Genotyping tools for G. duodenalis

Genotyping tools have been widely used in studies of the transmission of G. duodenalis in humans and animals. This has led to the identification of eight genotypes or assemblages of G. duodenalis that differ from each other in host range. Among them, assemblages A and B are major human-pathogens, but are also found in various mammals. In contrast, assemblages C to H are host-adapted G. duodenalis genotypes, being found in specific groups of animals (Table 2). Therefore, assemblages C and D are mostly found in canine animals, assemblage E in hoofed livestock and wildlife, assemblage F in cats, assemblage G in rodents, and assemblage H in seals and other marine mammals.

Early genotyping of G. duodenalis had used mostly PCR-sequence analysis of the SSU rRNA gene. Because of the low sequence polymorphism at this genetic locus, the use of these tools have led to the erroneous detections of G. duodenalis assemblages C, D, E and F in a small number of human specimens. More recent genotyping assays employ PCR targeting the more polymorphic β-giardin (bg), glutamate dehydrogenase (gdh), and triosephosphate isomerase (tpi) genes. Because differences in the sensitivity of the PCR assays and occasional discrepant genotyping results among genetic loci, it is recommended that G. duodenalis genotypes be determined by multilocus analysis, although human specimens genotyped at the three markers occasionally cannot be assigned unequivocally to either assemblage A or B.

DNA sequence analysis of PCR products are generally used in determination of G. duodenalis genotypes. However, RFLP analysis of PCR products is still used in resource-limited countries. This approach needs validation, as intragenotypic nucleotide variations in these polymorphic genes are common and are generally dispersed across the entire genes, leading to possible variations in RFLP patterns within some G. duodenalis assemblages such as assemblage B. In recent years, assemblage-specific PCR and qPCR assays have been developed for the detection of G. duodenalis assemblages A, B and E in clinical specimens. These assays are valuable in the detection of concurrent infections with mixed G. duodenalis genotypes.

The use of genotyping tools have identified mostly the presence of G. duodenalis assemblages A and B in humans. Globally, more humans are infected with assemblage B than assemblage A. This is especially the case for Asian, Oceanian, European, and African countries. Both genotypes appear to be equally distributed in the Americas and human infections in Mexico are largely caused by assemblage A.

Other G. duodenalis genotypes have been only occasionally detected in humans, with the possible exception of assemblage E, which has recently been detected in substantial numbers of human cases in rural areas of Egypt, Rio de Janeiro, Brazil, and Queensland, Australia. In contrast, animals are mostly infected with host-adapted G. duodenalis assemblages (C and D in canine animals, E in hoofed livestock, F in cats, and G in rodents) and assemblage A, with noticeable exception of nonhuman primates, beavers and a few other animals, which are commonly infected with assemblage B. The different distribution of G. duodenalis assemblages between humans and animals indicated that unlike in cryptosporidiosis, zoonotic transmission might not be important in giardiasis epidemiology.

Epidemiological studies of giardiasis in humans using genotyping tools have also revealed possible virulence differences between G. duodenalis assemblages A and B in humans. Some reports suggest that assemblage A is more virulent than assemblage B, while others, especially several recent ones, have indicated the opposite. One factor not considered was the intra-assemblage variation, which could possibly account for some differences between the studies. Data on genotype distribution of G. duodenalis in outbreaks of human giardiasis are helpful in delineating differences in virulence between assemblages A and B in humans, but only a dozen of outbreaks have been investigated thus far using genotyping tools. Most earlier reported giardiasis outbreaks were caused by assemblage B, with the exception of one waterborne outbreak in Canada and one hospital-associated outbreak in China, which were caused by assemblage A, indicating that assemblage B could be more virulent than assemblage A. A recent report, however, has shown that all four drinking water-associated outbreaks of giardiasis in small towns, British Columbia investigated were caused by mixed assemblages A and B, with B dominating. Despite the higher proportion of human infections due to assemblage B, assemblage A, especially the anthroponotic sub-assemblage AII, appears to be the dominant G. duodenalis in urban wastewater. Thus, in addition to possible differences in virulence, there could be other biological differences between the two major human-pathogenic G. duodenalis assemblages.

Subtyping tools for Cryptosporidium spp.

Subtyping tools have been used extensively in assessing intra-species diversity of C. parvum, C. hominis and other human-pathogenic Cryptosporidium spp. in endemic areas, role of zoonotic infections in cryptosporidiosis epidemiology, and contamination or infection sources during cryptosporidiosis outbreaks. Most subtyping tools are based on sequence analysis of the gene encoding the 60 kDa glycoprotein (gp60), which is known to be involved in invasion by Cryptosporidium spp.. The gp60 gene is highly polymorphic, categorizing C. parvum and C. hominis each into multiple subtype families by nucleotide sequence differences. Within each subtype family, subtypes differ from each other mostly in the number of tri-nucleotide repeats (TCA, TCG or TCT) region downstream of the signal peptide sequence of the gene.

A nomenclature system has been established for naming Cryptosporidium subtypes. The name of Cryptosporidium subtypes starts with the subtype family designation (Ia, Ib, Id, Ie, If, etc. for C. hominis; IIa, IIb, IIc, IId, etc. for C. parvum; IIIa, IIIb, IIIc, IIId, etc. for C. meleagridis). This is followed by the number of TCA (represented by the letter A), TCG (represented by the letter G), or TCT (represented by the letter T) repeats. For example, the subtype name IeA11G3T3 indicates that isolate belongs to the C. hominis subtype family Ie and has 11 copies of the TCA repeat, 3 copies of the TCG repeat, and 3 copies of the TCT repeat in the trinucleotide repeat region of the gene. In some subtype families, isolates may have identical number of the trinucleotide repeats, but differ from each other in the copy number of other repetitive sequences (designed as R at the end of the subtype name). For example, in the C. parvum IIa subtype family, subtypes have 1–3 copies of the ACATCA sequence immediately downstream from the trinucleotide repeats, which are represented by R1, R2 and R3. Similarly, subtypes within the C. hominis subtype families Ia and If frequently have different copies of a 15-bp repetitive sequence (AAGACGGTGGTAAGG or its variations in the Ia subtype family and AAGAAGGCAAAGAAG or its variations in the If subtype family). In addition, subtypes in the C. parvum subtype family IIc all have five copies of TCA and three copies of TCG repeats (IIcA5G3), but differ from each other in sequences of the 3′ region of the gene. These subtypes are differentiated from each by alphabetical extensions, such as IIcA5G3a and IIcA5G3b. The GenBank accession numbers of known Cryptosporidium subtype families are shown in Table 3.

Subtyping tools targeting the gp60 gene has been developed recently for several other human-pathogenic Cryptosporidium species such as C. meleagridis, C. ubiquitum, C. viatorum, and Cryptosporidium skunk genotype and chipmunk genotype I. These new subtyping tools are needed as PCR primers based on C. parvum and C. hominis sequences cannot amplify efficiently the gp60 gene of these divergent Cryptosporidium spp. Recent whole genome sequencing has greatly facilitated the design of PCR primers for them. Subtyping tools for other important zoonotic Cryptosporidium species such as C. felis and C. canis are not yet available due to the lack of whole genome sequence data.

There are significant phenotypic differences among subtype families of Cryptosporidium spp. Among the three most common C. parvum subtype families, IIa is mostly found in cattle, IId is mostly found in sheep and goats, whereas IIc is mostly found in humans, with the former two responsible for zoonotic cryptosporidiosis. However, in areas with extensive transmission of C. parvum in cattle, such as European countries, IIa subtypes can be found in lambs and goat kids, while in China, where IIa subtypes are largely absent in the country, dairy calves are mostly infected with IId subtypes. Host adaptation has also been identified in C. ubiquitum and C. tyzzeri based on sequence analysis of the gp60 gene. In humans, differences in clinical presentations and virulence have been observed among some C. hominis or C. parvum subtype families. At the subtype level, gp60 subtypes IbA10G2 of C. hominis and IIaA15G2R1 of C. parvum are widely distributed and responsible for numerous outbreaks of human cryptosporidiosis, probably due to their biological fitness and high virulence. In Peru, this subtype is associated with more clinical symptoms in infected children than other C. hominis subtypes.

Subtyping tools for G. duodenalis

Subtyping of G. duodenalis is currently done by sequence analysis of the PCR products of tpi, gdh, and bg genes, which leads to the identification of subtypes in addition to assemblages, based on the presence of SNPs within each assemblage. Within assemblage A, there are several subtypes at each of the genetic loci, frequently named as A1–A6. At the gdh locus, A1 and A5 subtypes have similar sequences, forming a subgroup that is divergent from the second subgroup formed by A2-A4. In contrast, the A6 subtype has very divergent sequence from other subtypes. The separation of the former two subgroups is not as evident at the tpi and bg loci, although the A6 subtype has very divergent sequences at both loci. There is apparent host adaption at the subtype level for the three common assemblage A subtypes at each genetic locus, with A1 subtype mostly found in animals, A2 mostly in humans, and A6 almost exclusively in wild ruminants.

Extensive genetic heterogeneity also occurs in assemblage B and other G. duodenalis genotypes. There are many more subtypes of assemblage B at these three genetic loci than assemblage A subtypes, but there are no robust formations of subgroups of assemblage B subtypes in phylogenetic analysis of tpi, gdh, and bg sequences. Indeed because of the presence of allelic sequence heterozygosity in assemblage B at these genotyping and subtyping loci, it may be difficult to subtype assemblage B specimens based on sequence analysis of PCR products from these loci. Many subtypes are also seen in assemblage E, with no formation of host-specific subgroups in nucleotide sequences from the three genetic loci. Subtyping of assemblage E specimens has been used extensively in recent studies of G. duodenalis transmission in ruminants in China.

Multilocus typing tools for Cryptosporidium spp.

Because of the likely occurrence of genetic recombination among isolates, results of gp60-based subtyping alone are unlikely to reflect intra-species diversity of Cryptosporidium spp. in disease endemic areas. To increase the resolution of gp60-based subtyping and facilitate population genetic studies, multilocus fragment (MLFT) or sequence (MLST) typing tools have been developed for C. parvum and C. hominis. These tools largely target sequences with microsatellite and minisatellite repeats, which are also known as variable-number tandem-repeats.

Results of isolate characterizations using MLFT tools have confirmed the existence of host-segregated C. parvum populations and identified differences in population genetic structure between C. parvum and C. hominis. In addition, significant geographic segregation of C. parvum populations has been identified among European countries, with population isolation by physical distance. However, because of a predominantly panmictic population structure, there has been no apparent geographic sub-structuring in C. parvum within Scotland and the Upper Midwest United States. As a result, distinct C. parvum MLFT types are frequently confined to individual farms. In Spain, C. parvum in cattle has a panmictic structure while C. parvum in sheep has a clonal structure, probably a reflection of differences in population genetic structure between gp60 IIa and IId subtype families. Recently, efforts have been made to evaluate existing MLFT markers and identify additional markers for robust MLFT analysis of C. parvum specimens.

MLST tools are increasingly used in genetic characterizations of C. parvum and C. hominis specimens from humans and animals. Compared with MLFT, MLST tools have the advantage of unambiguous comparability of nucleotide sequence data among distant laboratories and possible interpretation of MLST data in the context of whole genome sequence data (for example, it is possible to infer MLST types based on whole genome sequences for comparative analysis of existing data). The use of these tools in the characterizations of specimens from endemic areas has identified mostly a clonal population structure for C. hominis and the existence of geographically segregated C. hominis subpopulations among developing countries, with populations in Jamaica and Peru significantly different from those in India and Kenya. It has further supported the clonal nature of C. parvum IId subtypes from diverse areas. A systematic MLST analysis of 32 polymorphic markers in chromosome 6 had shown a high genetic heterogeneity within C. hominis IbA10G2 specimens from children living in a small community in Lima, Peru. Despite the overall clonal population structure of C. hominis subtypes in the community, genetic recombination was detected in the virulent IbA10G2 subtype. Genetic recombination was also shown to be a driving force for the emergence of the major C. hominis outbreak subtype IaA28R4 and C. parvum outbreak subtype IIaA15G2R1 in the United States.

One of the MLST tools has been used effectively in the identification of a clonal population structure of C. meleagridis and the likely occurrence of cross-species transmission of infections between birds and human in Lima, Peru. Based on some of the same targets and additional ones from comparative genomic analysis, a MLST tool has been developed for C. ubiquitum, which has confirmed the existence of host-adapted C. ubiquitum populations and likely occurrence of genetic recombination among rodent-adapted gp60 subtype families. An MLST tool has also been developed for the gastric species C. andersoni and C. muris. It has been used effectively in population genetic characterizations of bovine C. andersoni in China, with the identification of mostly an epidemic population structure.

Multilocus typing tools for G. duodenalis

There are currently no high-resolution MLST tools for G. duodenalis. The multilocus genotyping tool based on sequence analysis of the gdh, bg, and tpi loci has been used as a MLST tool in subtyping assemblages A, B and E as discussed above. Data from these loci have even been used in population genetic characterizations of the pathogen in humans and animals. An MLST subtype nomenclature system has been developed for assemblage A based on sequence analysis of the bg, gdh, and tpi genes. This has allowed the identification of three sub-assemblages within assemblage A: AI, AII, and AIII (Table 4). Similar to the existence of host-adaptation at the subtype level, there are differences in the distribution of these sub-assemblages among hosts, with AI dominating in animals, AII dominating in humans, and AIII mostly in wild ruminants, although the former two are both human pathogens.

In contrast, sequence analysis of assemblage B specimens at the three genetic loci has identified numerous subtypes, which are inconsistently segregated among loci. Thus, the BIII and BIV sub-assemblages originally identified by allozyme analysis are not supported by MLST analysis. MLST analysis of assemblage B specimens is further complicated by allelic sequence heterozygosity at some of the genetic loci. This also occurs in assemblages C, D and E, but not assemblage A at these genotyping loci. Nevertheless, geographic segregation and host-adaptation have been identified in assemblage B in captive nonhuman primates in China using the current MLST tool; isolates from China were significantly different from those from Italy and Sweden, and isolates from ring-tailed lemurs and squirrel monkeys were genetically different from those from other nonhuman primates. Clearly, better MLST tools targeting genetic loci with homogenous sequences are needed for high-resolution typing and population genetic characterizations of G. duodenalis. This becomes feasible with the recent identification of numerous highly polymorphic genes within assemblage A or B through comparative genomics.

Comparative genomic analysis of Cryptosporidium spp.

Comparative genomics is increasingly used in the characterization of human pathogenic Cryptosporidium spp. For a considerable period, whole genome sequence (WGS) data are available from only four laboratory-propagated Cryptosporidium isolates. With the recent development of next generation sequencing techniques and procedures for isolation and enrichment of pure Cryptosporidium DNA, many Cryptosporidium isolates have been sequenced and data of them are available in public databases such as NCBI and CryptoDB. As expected, most of the isolates sequenced are from C. parvum and C. hominis, but several other Cryptosporidium species have also been sequenced, including C. meleagridis, C. ubiquitum, C. baileyi, and C. andersoni.

Despite the high sequence similarity (~ 97%) between C. parvum and C. hominis genomes, comparative analyses of the WGS data have identified some differences in gene content between the two major human-pathogenic Cryptosporidium species. The gene gains and losses mainly occur in the subtelomeric regions of several chromosomes, and involve mostly two Cryptosporidium-specific secretory protein families: MEDLE proteins and insulinase-like proteases. These differences have also been observed between them and C. ubiquitum, and between C. parvum gp60 subtype families IIa and IId. In addition to these gene gains and losses, these Cryptosporidium species and C. parvum have highly divergent sequences in genes encoding invasion-associated and immunodominant mucin proteins and other families of secretory proteins. These genetic differences could be responsible for some of the phenotypic differences among closely related Cryptosporidium species and host-adapted C. parvum subtype families. This has been recently supported by a preliminary study of MEDLE-2, which was shown to be involved in the invasion of host cells.

Comparative genomic analysis of C. hominis isolates from the United States has confirmed the occurrence of genetic recombination in virulent outbreak subtypes IbA10G2 and IaA28R4. Genomes of the two subtypes are highly similar, with major sequence differences occurring at the 5′ (around cgd6_60) and 3′ (around cgd6_5270) subtelomeric regions and around gp60 (cgd6_1080), all within chromosome 6. In these areas, the genomes have shown some hallmarks of genetic recombination, with bi-allelic sequences in these areas and a mosaic pattern in sequence types across the chromosome. As the sequence differences involve a secretory protease (cgd6_60) and an invasion-related mucin protein (gp60), both of which are among the most polymorphic genes in Cryptosporidium spp., genetic recombination could play a potential role in the emergence of hyper-transmissible C. hominis subtypes, as previously indicated by MLST characterizations of IbA10G2 in Peru and IaA28R4 in the United States.

The genomes of C. hominis IbA10G2 in European countries appear to be highly conserved. A comparative genomic analysis of WGS data from several IbA10G2 isolates has identified fewer than 50 SNPs among most isolates from Europe, with the exception of one isolate (SWEH9) from Spain. In contrast, the genome of IbA10G2 isolates from Guatemala is divergent from genomes of European isolates. This is in contrast with IbA10G2 in the United States, where two types of genomes are present among the few isolates sequenced, one of which is similar to the one in European countries. Several SNP features unique to the C. hominis IbA10G2 subtype have been identified in a recent comparative genomic analysis, including the restoration of a loss-of-function mutation seen in other C. hominis subtypes from developing countries. The high genome sequence conservation in IbA10G2 presents a significant challenge in epidemiologic tracking of this subtype, which is a major outbreak subtype in European countries and responsible for almost all autochthonous C. hominis infections there. Recently, based on WGS data from 17 isolates, a panels of nine loci with 10 SNVs were selected for the development of a PCR-next generation sequencing technique, which had identified 10 sequence types among 44 clinical isolates in Sweden.

Comparative genomic analysis of G. duodenalis

WGS data are available from several laboratory isolates of assemblages A, B and E. Results of comparative analysis have shown significant differences among the three genotypes in genome size, gene content, gene arrangement, nucleotide sequences, and surface molecule repertoires. For example, the genomes of AI and B have only about 70% nucleotide sequence identity across the syntenic regions. The divergence between sub-assemblages AI and AII is approximately 1% in sequences, with differences mainly localized to the variable regions of the genomes. There are approximately 35 AII-specific genes but only two AI-specific genes.

An approach has been developed recently for whole genome sequencing of G. duodenalis in clinical specimens. It uses a procedure similar to the one developed for Cryptosporidium spp. for isolating G. duodenalis cysts in clinical specimens. This could facilitate WGS-based characterization of G. duodenalis specimens and increase the use of comparative genomics in the identification of contamination source during giardiasis outbreaks, as indicated by the retrospective investigation of a drinking water-associated outbreak in Creston, British Columbia, Canada.

Use of molecular diagnostic tools in epidemiological studies of Cryptosporidium spp. and G. duodenalis in USA

The use of molecular diagnostic tools in epidemiologic studies has significantly improved our understanding of the transmission of waterborne protozoan parasites in the United States, especially Cryptosporidium spp. Compared with other industrialized nations, the United States has a high diversity of Cryptosporidium spp. in humans at both species and subtype levels. In addition to C. hominis and C. parvum, several emerging Cryptosporidium species or genotypes, such as C. ubiquitum and chipmunk genotype I, are responsible for substantial numbers of cryptosporidiosis cases, especially in rural states. For C. ubiquitum, humans in the United States are infected with rodent-adapted subtype families XIIb–XIId, whereas those in the United Kingdom are mainly with ruminant-adapted subtype family XIIa. As C. ubiquitum is one of the most common Cryptosporidium species in drinking source water in the United States and rodent-adapted subtype families are also the dominant C. ubiquitum subtypes in drinking source water, consumption of drinking water contaminated by infected wildlife could be the source of human infections. The genetic similarity in Cryptosporidium chipmunk genotype I among isolates from humans, wild mammals, and drinking source water supports the potential role of wildlife and drinking source water in the transmission of emerging Cryptosporidium spp. in rural United States.

Genotyping and subtyping data on C. parvum occurrence in sporadic cases of cryptosporidiosis indicate that farm animals, especially pre-weaned calves, are important in cryptosporidiosis epidemiology in rural United States. Like in the United Kingdom, there are apparent differences in the distribution of C. hominis and C. parvum between rural states and other states, with C. parvum as the dominant species in sporadic cases in Minnesota and Wisconsin and C. hominis as the dominant species in southern states. A high C. parvum subtype diversity was identified in humans in Minnesota and Wisconsin. As the dominant subtypes are also common ones in pre-weaned calves in the United States, these animals are potential sources of zoonotic infections in humans. This is also supported by the numerous reports of calf-contact associated outbreaks of human cryptosporidiosis in the United States and results of case-control studies, and by results of MLFT analysis of C. parvum isolates from humans and cattle in Upper Midwest states. The recent identification of cryptosporidiosis occurrence in Nebraska due to occupational animal exposure, especially to cattle, reinforces the importance of zoonotic infection in cryptosporidiosis epidemiology in rural U.S. states.

Among the > 20 C. parvum subtypes found in humans in the United States, IIaA15G2R1, is the dominant C. parvum subtype, responsible for numerous outbreaks of cryptosporidiosis (Table 5). Data from a recent MLST study have shown extensive heterogeneity within IIaA15G2R1. Because of the widespread occurrence of genetic recombination within the subtype, there is frequent discordance in subtyping results between gp60 and other genetic markers. As a result, there is no clear population diversion between IIaA15G2R and non-IIaA15G2R subtypes. Other population differentiation exists within C. parvum, evident by the presence of one subpopulation with an epidemic population structure and wide geographic distribution and another subpopulation with a clonal population structure and geographic segregation. Genetic recombination between the epidemic and geographically segregated C. parvum populations appears to be associated with the emergence of the hyper-transmissible IIaA15G2R1 subtype, which is also the dominant C. parvum subtype in humans in other industrialized nations. Thus, the IIaA15G2R1 subtype at the gp60 locus is probably a fitness marker for C. parvum and the wide spread of IIaA15G2R1 subtype around the world appears independent of sequence characteristics at other genetic loci. This needs confirmations from comparative genomic analysis of WGS data generated in recent years.

A high genetic heterogeneity is also present in C. hominis in the United States. In addition to the virulent IbA10G2 subtype, which is the dominant C. hominis subtype in European countries and Australia, several other C. hominis subtypes are common in sporadic and outbreaks cases in the United States, such as IaA28R4 and IfA12G1R5 (Table 5). In early years, IbA10G2 was the dominant C. hominis subtype in the United States. A new subtype, IaA28R4, however, appeared in the United States in 2005 and by 2007 it had become the dominant C. hominis subtype for sporadic cases and outbreaks in the United States (Table 5). Accompanied with the occurrence of the new subtype was a near three-fold increase in the incidence of cryptosporidiosis in the United States during 2004 to 2008. In recent years, subtype IaA28R4 has been replaced by IfA12G1R5, which was first seen in AIDS patients and was responsible for a 2009 cryptosporidiosis outbreak associated with the care of an AIDS patient in Oregon. Since 2013, it has emerged as the dominant C. hominis subtype in sporadic and outbreak-associated cases. During this period, the incidence of cryptosporidiosis in the United States has remained at a high level.

Recent MLST and comparative genomic analysis indicate that genetic recombination has played a major role in the emergence of hyper-transmissible C. hominis subtypes in the United States. Results of the MLST analysis indicate that there were at least two origins of IaA28R4 within the United States. Like IIaA15G2R1 within the United States, discordant subtyping results of IaA28R4 isolates were observed between gp60 and other genetic markers, indicating that the emergence of these IaA28R4 variants was largely due to genetic recombination. Although IaA28R4 was first identified in a cryptosporidiosis outbreak in Ohio in 2005 and was the cause of another outbreak in the same area in 2008, IaA28R4 circulating in Ohio was genetically different from IaA28R4 in southwestern states. Results of comparative genomic analysis indicate that the genomes of two IaA28R4 isolates were very similar to those of IbA10G2, indicating that IaA28R4 could be the progeny of genetic recombination of IbA10G2 and another C. hominis subtype of the Ia subtype family. The genetic recombination was most obvious in the subtelomeric and gp60 regions of chromosome 6. Thus, genetic recombination might lead to the emergence of hyper-transmissible gp60 subtypes of both C. parvum and C. hominis.

Molecular characterizations of animal and human isolates have also played a role in clarifications of the potential role of beavers in the transmission of G. duodenalis in the United States. Giardiasis has been popularly known as beaver fever in North America, probably because some hikers and campers became ill from drinking stream water presumably contaminated with Giardia cysts from beavers. This was supported by results of investigations of several waterborne outbreaks of giardiasis in late 1970s and early 1980s, in which beavers in drinking source watershed were found to be shedding G. duodenalis cysts. Results of molecular characterizations have shown that beavers are frequently infected with G. duodenalis assemblage B, which is the major cause of giardiasis outbreaks in the United States. In studies conducted in Maryland and Massachusetts, 11 beavers were identified having assemblage B. In one drinking water-associated outbreak of giardiasis in New Hampshire, the assemblage B subtype found in human cases was different from known subtypes in humans, but identical to the one previously found in beavers in Massachusetts. Although these reports did not provide any concrete evidence of the initial contamination source, they suggest that beavers could be environmental reservoirs for sustaining the contamination of drinking source water by G. duodenalis assemblage B. This has been supported by the investigation of one waterborne giardiasis outbreak in Creston, British Columbia, in which one beaver was identified by epidemiologic investigations as the likely cause of source water contamination. WGS analysis of one beaver isolate and one human isolate from the outbreak generated very similar genomes of sub-assemblage AI.

Conclusions and perspectives

The use of molecular diagnostic tools has significantly improved our understanding of cryptosporidiosis epidemiology. It has led to the identification of major differences in infection sources of Cryptosporidium spp. in humans between developing countries and industrialized nations, differences in clinical presentations and virulence among Cryptosporidium species and C. hominis subtypes, and roles of genetic recombination in the emergence of virulent and hyper-transmissible C. hominis and C. parvum subtypes. These tools are now widely used in outbreak investigations and surveillance of cryptosporidiosis in industrialized nations. With the introduction of whole genome sequencing and comparative genomics in Cryptosporidium characterization, it is expected that molecular epidemiology of cryptosporidiosis would remain a highly active research area.

Despite these major achievements, many challenges remain. After over 20 years of major investments in drinking water treatment as a response to the massive 1993 Milwaukee outbreak, cryptosporidiosis remains the most important waterborne disease in industrialized nations, incidence of cryptosporidiosis in the United States has increased nearly threefold since 2005, and drinking water-associated outbreaks of cryptosporidiosis still present a significant public health and economic problem in European countries. The identification of environmental and epidemiological factors involved requires systematic molecular characterizations of Cryptosporidium specimens submitted to national cryptosporidiosis surveillance systems. The latter has been in place for some time in the United Kingdom but is in the early stage of implementation in the United States as part of the CryptoNet program. While genotype and subtype data are useful in the identification and investigation of zoonotic outbreaks due to unusual C. parvum subtypes and differentiation of autochthonous and travel-associated C. hominis infections in European countries, their utilities are limited in molecular surveillance of the dominant C. hominis subtypes in Europe (IbA10G2) and the United States (currently IfA12G1R5). This was demonstrated in the investigation of an increased transmission of IbA10G2 in multiple European countries in late summer, 2012, in which molecular characterizations did not identify an endemic source. In fact, advanced molecular analysis beyond gp60 sequence analysis would not increase significantly the discrimination of these hyper-transmissible C. hominis subtypes, including comparative genomic analysis. Supplemental epidemiological data are helpful in the investigation of such outbreaks.

Molecular analysis probably can exert its biggest impact on improving knowledge of cryptosporidiosis epidemiology in developing countries, where cryptosporidiosis has been identified as the second most cause of modest to severe diarrhea and diarrhea-associated mortality in young children, and diverse C. hominis subtypes are circulating in endemic areas. While data from genotyping and subtyping studies of clinical specimens have suggested that anthroponotic infection is important in cryptosporidiosis epidemiology in these areas, the role of various transmission routes is not very clear for the formulation of evidence-based intervention strategies. This would require the integrated use of molecular diagnostic tools in well-designed epidemiological studies, which usually involve the use of longitudinal cohort or case-control design, or regression and multivariate analyses of risk factors. Thus far, such studies are few in numbers, and WGS and comparative genomic analyses have not been used in characterizations of Cryptosporidium spp. in developing countries. The population genetics of C. hominis in developing countries is probably complicated; genetic recombination among diverse subtypes could occur more frequently as a result of high cryptosporidiosis endemicity. Advanced molecular characterizations of specimens from well-designed epidemiologic investigations will likely lead to better understanding of the spread of virulent C. hominis subtypes in under-developed communities, as demonstrated in a MLST characterization of C. hominis from a longitudinal cohort study of cryptosporidiosis conducted in a shantytown in Lima, Peru.

In contrast, molecular analysis of G. duodenalis has thus far mostly generated data useful in understanding of zoonotic potential of isolates in various animals. Because of host-adaptation at both genotype and subtype levels, zoonotic transmission of G. duodenalis is likely to be less important in giardiasis epidemiology compared with cryptosporidiosis. Much of the current attentions on zoonotic transmission of G. duodenalis have focused on assemblage A because of its wide occurrence in animals compared with assemblage B. As humans are more commonly infected with assemblage B and there is a difference in the distribution of assemblage A subtypes between humans and animals, more attention should be directed to zoonotic transmission of assemblage B, which appears to be to the dominant G. duodenalis genotype in a few groups of animals, such as beavers, muskrats, chinchillas, rabbits, horses, and nonhuman primates. Some of the animals are aquatic or their fecal materials can be easily transported to water, thus could contribute to source water contamination with G. duodenalis assemblage B. The identification of assemblage E humans in several recent studies indicates that it could be an emerging human pathogenic G. duodenalis genotype, even in urban areas. Additional molecular epidemiological investigations should be conducted to identify its source and transmission routes.

The use of molecular diagnostic tools has unfortunately not improved significantly our knowledge of endemic and epidemic giardiasis in humans. This is due to both technical difficulties and neglect. Thus far, most molecular epidemiological studies of giardiasis in humans are conducted with clinical specimens from outpatients. The long duration of cyst shedding after the initial infection, presence of allelic sequence heterozygosity in some genotyping and subtyping markers, high occurrence of concurrent infections with mixed genotypes, and multiple episodes of infections in early childhood have made it challenging to delineate the transmission mechanism and virulence of G. duodenalis in humans. With recent developments in whole genome sequencing and comparative genomic analysis of G. duodenalis, new genotyping and subtyping markers can be identified for the development of next generation molecular tools. As assemblages A and B evolve very differently and have divergent genomes, it may be useful to develop assemblage-specific subtyping tools. The use of these advanced molecular diagnostic tools in epidemiological investigations of endemic giardiasis in developing countries and in surveillance and outbreak investigations of epidemic giardiasis in industrialized nations, as performed in several recent studies, may lead to improved understanding of molecular epidemiology of giardiasis.

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    Efstratiou A., Ongerth J.E., Karanis P. Waterborne transmission of protozoan parasites: review of worldwide outbreaks - an update 2011–2016. Water Res. 2017;114:14-22.

    Not in our library10.1016/j.watres.2017.01.036 (opens in a new tab)

  55. 55.

    Eibach D., Krumkamp R., Al-Emran H.M., Sarpong N., Hagen R.M., Adu-Sarkodie Y., Tannich E., May J. Molecular characterization of Cryptosporidium spp. among children in rural Ghana. PLoS Negl. Trop. Dis. 2015;9.

    Not in our library10.1371/journal.pntd.0003551 (opens in a new tab)

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    El Basha N.R., Zaki M.M., Hassanin O.M., Rehan M.K., Omran D. Giardia assemblages A and B in diarrheic patients: a comparative study in Egyptian children and adults. J. Parasitol. 2016;102:69-74.

    Not in our library10.1645/14-676 (opens in a new tab)

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    Elwin K., Hadfield S.J., Robinson G., Chalmers R.M. The epidemiology of sporadic human infections with unusual cryptosporidia detected during routine typing in England and Wales, 2000-2008. Epidemiol. Infect. 2012;140:673-683.

    Not in our library10.1017/s0950268811000860 (opens in a new tab)

  58. 58.

    Elwin K., Hadfield S.J., Robinson G., Crouch N.D., Chalmers R.M. Cryptosporidium viatorum n. sp. (Apicomplexa: Cryptosporidiidae) among travellers returning to Great Britain from the Indian subcontinent, 2007–2011. Int. J. Parasitol. 2012;42:675-682.

    Not in our library10.1016/j.ijpara.2012.04.016 (opens in a new tab)

  59. 59.

    Emery S.J., van Sluyter S., Haynes P.A. Proteomic analysis in Giardia duodenalis yields insights into strain virulence and antigenic variation. Proteomics. 2014;14:2523-2534.

    Not in our library10.1002/pmic.201400144 (opens in a new tab)

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    Fahmy H.M., El-Serougi A.O., El Deeb H.K., Hussein H.M., Abou-Seri H.M., Klotz C., Aebischer T., El Sayed Khalifa Mohamed K. Giardia duodenalis assemblages in Egyptian children with diarrhea. Eur. J. Clin. Microbiol. Infect. Dis. 2015;34:1573-1581.

    Not in our library10.1007/s10096-015-2389-7 (opens in a new tab)

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    Fantinatti M., Bello A.R., Fernandes O., Da-Cruz A.M. Identification of Giardia lamblia assemblage E in humans points to a new anthropozoonotic cycle. J Infect Dis. 2016;214:1256-1259.

    Not in our library10.1093/infdis/jiw361 (opens in a new tab)

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    Faria C.P., Zanini G.M., Dias G.S., da Silva S., Sousa Mdo C. Molecular characterization of Giardia lamblia: first report of assemblage B in human isolates from Rio de Janeiro (Brazil). PLoS ONE. 2016;11.

    Not in our library10.1371/journal.pone.0160762 (opens in a new tab)

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    Faria C.P., Zanini G.M., Dias G.S., da Silva S., Sousa M.D.C. New multilocus genotypes of Giardia lamblia human isolates. Infect. Genet. Evol. 2017;54:128-137.

    Not in our library10.1016/j.meegid.2017.06.028 (opens in a new tab)

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    Faria C.P., Zanini G.M., Dias G.S., Sousa M.D.C. Associations of Giardia lamblia assemblages with HIV infections and symptomatology: HIV virus and assemblage B were they born to each other? Acta Trop. 2017;172:80-85.

    Not in our library10.1016/j.actatropica.2017.04.026 (opens in a new tab)

  65. 65.

    Faubert G.M. Evidence that giardiasis is a zoonosis. Parasitol. Today. 1988;4:66-68.

    Not in our library10.1016/0169-4758(88)90197-4 (opens in a new tab)

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    Fayer R., Santin M., Trout J.M., DeStefano S., Koenen K., Kaur T. Prevalence of Microsporidia, Cryptosporidium spp., and Giardia spp. in beavers (Castor canadensis) in Massachusetts. J. Zoo Wildl. Med. 2006;37:492-497.

    Not in our library10.1638/06-013.1 (opens in a new tab)

  67. 67.

    Fayer R., Santin M., Macarisin D. Cryptosporidium ubiquitum n. sp. in animals and humans. Vet. Parasitol. 2010;172:23-32.

    Not in our library10.1016/j.vetpar.2010.04.028 (opens in a new tab)

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    Feltus D.C., Giddings C.W., Schneck B.L., Monson T., Warshauer D., McEvoy J.M. Evidence supporting zoonotic transmission of Cryptosporidium in Wisconsin. J. Clin. Microbiol. 2006;44:4303-4308.

    Not in our library10.1128/jcm.01067-06 (opens in a new tab)

  69. 69.

    Feng Y., Xiao L. Zoonotic potential and molecular epidemiology of Giardia species and giardiasis. Clin. Microbiol. Rev. 2011;24:110-140.

    Not in our library10.1128/cmr.00033-10 (opens in a new tab)

  70. 70.

    Feng Y., Ortega Y., He G., Das P., Xu M., Zhang X., Fayer R., Gatei W., Cama V., Xiao L. Wide geographic distribution of Cryptosporidium bovis and the deer-like genotype in bovines. Vet. Parasitol. 2007;144:1-9.

    Not in our library10.1016/j.vetpar.2006.10.001 (opens in a new tab)

  71. 71.

    Feng Y., Lal A.A., Li N., Xiao L. Subtypes of Cryptosporidium spp. in mice and other small mammals. Exp. Parasitol. 2011;127:238-242.

    Not in our library10.1016/j.exppara.2010.08.002 (opens in a new tab)

  72. 72.

    Feng Y., Yang W., Ryan U., Zhang L., Kvac M., Koudela B., Modry D., Li N., Fayer R., Xiao L. Development of a multilocus sequence tool for typing Cryptosporidium muris and Cryptosporidium andersoni. J. Clin. Microbiol. 2011;49:34-41.

    Not in our library10.1128/jcm.01329-10 (opens in a new tab)

  73. 73.

    Feng Y., Wang L., Duan L., Gomez-Puerta L.A., Zhang L., Zhao X., Hu J., Zhang N., Xiao L. Extended outbreak of cryptosporidiosis in a pediatric hospital, China. Emerg. Infect. Dis. 2012;18:312-314.

    Not in our library10.3201/eid1802.110666 (opens in a new tab)

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    Feng Y., Torres E., Li N., Wang L., Bowman D., Xiao L. Population genetic characterisation of dominant Cryptosporidium parvum subtype IIaA15G2R1. Int. J. Parasitol. 2013;43:1141-1147.

    Not in our library10.1016/j.ijpara.2013.09.002 (opens in a new tab)

  75. 75.

    Feng Y., Tiao N., Li N., Hlavsa M., Xiao L. Multilocus sequence typing of an emerging Cryptosporidium hominis subtype in the United States. J. Clin. Microbiol. 2014;52:524-530.

    Not in our library10.1128/jcm.02973-13 (opens in a new tab)

  76. 76.

    Feng Y., Li N., Roellig D.M., Kelley A., Liu G., Amer S., Tang K., Zhang L., Xiao L. Comparative genomic analysis of the IId subtype family of Cryptosporidium parvum. Int. J. Parasitol. 2017;47:281-290.

    Not in our library10.1016/j.ijpara.2016.12.002 (opens in a new tab)

  77. 77.

    Figgatt M., Mergen K., Kimelstein D., Mahoney D.M., Newman A., Nicholas D., Ricupero K., Cafiero T., Corry D., Ade J., Kurpiel P., Madison-Antenucci S., Anand M. Giardiasis outbreak associated with asymptomatic food handlers in New York State, 2015. J. Food Prot. 2017;837-841.

    Not in our library10.4315/0362-028x.jfp-16-415 (opens in a new tab)

  78. 78.

    Foronda P., Bargues M.D., Abreu-Acosta N., Periago M.V., Valero M.A., Valladares B., Mas-Coma S. Identification of genotypes of Giardia intestinalis of human isolates in Egypt. Parasitol. Res. 2008;103:1177-1181.

    Not in our library10.1007/s00436-008-1113-2 (opens in a new tab)

  79. 79.

    Fournet N., Deege M.P., Urbanus A.T., Nichols G., Rosner B.M., Chalmers R.M., Gorton R., Pollock K.G., van der Giessen J.W., Wever P.C., Dorigo-Zetsma J.W., Mulder B., Mank T.G., Overdevest I., Kusters J.G., van Pelt W., Kortbeek L.M. Simultaneous increase of Cryptosporidium infections in the Netherlands, the United Kingdom and Germany in late summer season, 2012. Euro Surveill. 2013;18.

    Not in our library

  80. 80.
    Draft Genome Sequencing of Giardia intestinalis Assemblage B Isolate GS: Is Human Giardiasis Caused by Two Different Species? · 2009
    No notice
  81. 81.
    Estimates of global, regional, and national morbidity, mortality, and aetiologies of diarrhoeal diseases: a systematic analysis for the Global Burden of Disease Study 2015 · 2017
    Corrected
  82. 82.

    Gabin-Garcia L.B., Bartolome C., Abal-Fabeiro J.L., Mendez S., Llovo J., Maside X. Strong genetic structure revealed by multilocus patterns of variation in Giardia duodenalis isolates of patients from Galicia (NW-Iberian Peninsula). Infect. Genet. Evol. 2017;48:131-141.

    Not in our library10.1016/j.meegid.2016.12.014 (opens in a new tab)

  83. 83.

    Gatei W., Ashford R.W., Beeching N.J., Kamwati S.K., Greensill J., Hart C.A. Cryptosporidium muris infection in an HIV-infected adult, Kenya. Emerg. Infect. Dis. 2002;8:204-206.

    Not in our library10.3201/eid0802.010256 (opens in a new tab)

  84. 84.

    Gatei W., Greensill J., Ashford R.W., Cuevas L.E., Parry C.M., Cunliffe N.A., Beeching N.J., Hart C.A. Molecular analysis of the 18S rRNA gene of Cryptosporidium parasites from patients with or without human immunodeficiency virus infections living in Kenya, Malawi, Brazil, the United Kingdom, and Vietnam. J. Clin. Microbiol. 2003;41:1458-1462.

    Not in our library10.1128/jcm.41.4.1458-1462.2003 (opens in a new tab)

  85. 85.

    Gatei W., Hart C.A., Gilman R.H., Das P., Cama V., Xiao L. Development of a multilocus sequence typing tool for Cryptosporidium hominis. J. Eukaryot. Microbiol. 2006;53:S43-S48.

    Not in our library10.1111/j.1550-7408.2006.00169.x (opens in a new tab)

  86. 86.

    Gatei W., Wamae C.N., Mbae C., Waruru A., Mulinge E., Waithera T., Gatika S.M., Kamwati S.K., Revathi G., Hart C.A. Cryptosporidiosis: prevalence, genotype analysis, and symptoms associated with infections in children in Kenya. Am. J. Trop. Med. Hyg. 2006;75:78-82.

    Not in our library

  87. 87.

    Gatei W., Das P., Dutta P., Sen A., Cama V., Lal A.A., Xiao L. Multilocus sequence typing and genetic structure of Cryptosporidium hominis from children in Kolkata, India. Infect. Genet. Evol. 2007;7:197-205.

    Not in our library10.1016/j.meegid.2006.08.006 (opens in a new tab)

  88. 88.

    Gatei W., Barrett D., Lindo J.F., Eldemire-Shearer D., Cama V., Xiao L. Unique Cryptosporidium population in HIV-Infected persons, Jamaica. Emerg. Infect. Dis. 2008;14:841-843.

    Not in our library10.3201/eid1405.071277 (opens in a new tab)

  89. 89.

    Gelanew T., Lalle M., Hailu A., Pozio E., Caccio S.M. Molecular characterization of human isolates of Giardia duodenalis from Ethiopia. Acta Trop. 2007;102:92-99.

    Not in our library10.1016/j.actatropica.2007.04.003 (opens in a new tab)

  90. 90.

    Guo Y., Cebelinski E., Matusevich C., Alderisio K.A., Lebbad M., McEvoy J., Roellig D.M., Yang C., Feng Y., Xiao L. Subtyping novel zoonotic pathogen Cryptosporidium chipmunk genotype I. J. Clin. Microbiol. 2015;53:1648-1654.

    Not in our library10.1128/jcm.03436-14 (opens in a new tab)

  91. 91.

    Guo Y., Li N., Lysen C., Frace M., Tang K., Sammons S., Roellig D.M., Feng Y., Xiao L. Isolation and enrichment of Cryptosporidium DNA and verification of DNA purity for whole-genome sequencing. J. Clin. Microbiol. 2015;53:641-647.

    Not in our library10.1128/jcm.02962-14 (opens in a new tab)

  92. 92.

    Guo Y., Tang K., Rowe L.A., Li N., Roellig D.M., Knipe K., Frace M., Yang C., Feng Y., Xiao L. Comparative genomic analysis reveals occurrence of genetic recombination in virulent Cryptosporidium hominis subtypes and telomeric gene duplications in Cryptosporidium parvum. BMC Genomics. 2015;16:320.

    Not in our library10.1186/s12864-015-1517-1 (opens in a new tab)

  93. 93.

    Guyot K., Follet-Dumoulin A., Lelievre E., Sarfati C., Rabodonirina M., Nevez G., Cailliez J.C., Camus D., Dei-Cas E. Molecular characterization of Cryptosporidium isolates obtained from humans in France. J. Clin. Microbiol. 2001;39:3472-3480.

    Not in our library10.1128/jcm.39.10.3472-3480.2001 (opens in a new tab)

  94. 94.

    Hadfield S.J., Chalmers R.M. Detection and characterization of Cryptosporidium cuniculus by real-time PCR. Parasitol. Res. 2012;111:1385-1390.

    Not in our library10.1007/s00436-012-2874-1 (opens in a new tab)

  95. 95.

    Hadfield S.J., Robinson G., Elwin K., Chalmers R.M. Detection and differentiation of Cryptosporidium spp. in human clinical samples using real-time PCR. J. Clin. Microbiol. 2011;49:918-924.

    Not in our library10.1128/jcm.01733-10 (opens in a new tab)

  96. 96.

    Hadfield S.J., Pachebat J.A., Swain M.T., Robinson G., Cameron S.J., Alexander J., Hegarty M.J., Elwin K., Chalmers R.M. Generation of whole genome sequences of new Cryptosporidium hominis and Cryptosporidium parvum isolates directly from stool samples. BMC Genomics. 2015;16:650.

    Not in our library10.1186/s12864-015-1805-9 (opens in a new tab)

  97. 97.

    Hanevik K., Bakken R., Brattbakk H.R., Saghaug C.S., Langeland N. Whole genome sequencing of clinical isolates of Giardia lamblia. Clin. Microbiol. Infect. 2015;21(192):e191-193.

    Not in our library10.1016/j.cmi.2014.08.014 (opens in a new tab)

  98. 98.

    Haque R., Roy S., Kabir M., Stroup S.E., Mondal D., Houpt E.R. Giardia assemblage A infection and diarrhea in Bangladesh. J Infect Dis. 2005;192:2171-2173.

    Not in our library10.1086/498169 (opens in a new tab)

  99. 99.

    Hasajova A., Valencakova A., Malcekova B., Danisova O., Halan M., Goldova M., Sak B., Kvetonova D., Kvac M., Halanova M. Significantly higher occurrence of Cryptosporidium infection in Roma children compared with non-Roma children in Slovakia. Eur. J. Clin. Microbiol. Infect. Dis. 2014;33:1401-1406.

    Not in our library10.1007/s10096-014-2082-2 (opens in a new tab)

  100. 100.

    Hatam-Nahavandi K., Mohebali M., Mahvi A.H., Keshavarz H., Mirjalali H., Rezaei S., Meamar A.R., Rezaeian M. Subtype analysis of Giardia duodenalis isolates from municipal and domestic raw wastewaters in Iran. Environ. Sci. Pollut. Res. Int. 2017;24:12740-12747.

    Not in our library10.1007/s11356-016-6316-y (opens in a new tab)

  101. 101.

    Helmy Y.A., Krucken J., Nockler K., von Samson-Himmelstjerna G., Zessin K.H. Molecular epidemiology of Cryptosporidium in livestock animals and humans in the Ismailia province of Egypt. Vet. Parasitol. 2013;193:15-24.

    Not in our library10.1016/j.vetpar.2012.12.015 (opens in a new tab)

  102. 102.

    Helmy Y.A., Klotz C., Wilking H., Krucken J., Nockler K., Von Samson-Himmelstjerna G., Zessin K.H., Aebischer T. Epidemiology of Giardia duodenalis infection in ruminant livestock and children in the Ismailia province of Egypt: insights by genetic characterization. Parasit. Vectors. 2014;7:321.

    Not in our library10.1186/1756-3305-7-321 (opens in a new tab)

  103. 103.

    Herges G.R., Widmer G., Clark M.E., Khan E., Giddings C.W., Brewer M., McEvoy J.M. Evidence that Cryptosporidium parvum populations are panmictic and unstructured in the Upper Midwest of the United States. Appl. Environ. Microbiol. 2012;78:8096-8101.

    Not in our library10.1128/aem.02105-12 (opens in a new tab)

  104. 104.

    Hlavsa M.C., Roberts V.A., Kahler A.M., Hilborn E.D., Mecher T.R., Beach M.J., Wade T.J., Yoder J.S., Centers for Disease, C., Prevention. Outbreaks of illness associated with recreational water–United States, 2011–2012. MMWR Morb. Mortal. Wkly Rep. 2015;64:668-672.

    Not in our library

  105. 105.

    Hlavsa M.C., Roellig D.M., Seabolt M.H., Kahler A.M., Murphy J.L., McKitt T.K., Geeter E.F., Dawsey R., Davidson S.L., Kim T.N., Tucker T.H., Iverson S.A., Garrett B., Fowle N., Collins J., Epperson G., Zusy S., Weiss J.R., Komatsu K., Rodriguez E., Patterson J.G., Sunenshine R., Taylor B., Cibulskas K., Denny L., Omura K., Tsorin B., Fullerton K.E., Xiao L. Using molecular characterization to support investigations of aquatic facility-associated outbreaks of cryptosporidiosis - Alabama, Arizona, and Ohio, 2016. MMWR Morb. Mortal. Wkly Rep. 2017;66:493-497.

    Not in our library10.15585/mmwr.mm6619a2 (opens in a new tab)

  106. 106.

    Homan W.L., Mank T.G. Human giardiasis: genotype linked differences in clinical symptomatology. Int. J. Parasitol. 2001;31:822-826.

    Not in our library10.1016/s0020-7519(01)00183-7 (opens in a new tab)

  107. 107.

    Hotchkiss E.J., Gilray J.A., Brennan M.L., Christley R.M., Morrison L.J., Jonsson N.N., Innes E.A., Katzer F. Development of a framework for genotyping bovine-derived Cryptosporidium parvum, using a multilocus fragment typing tool. Parasit. Vectors. 2015;8:500.

    Not in our library10.1186/s13071-015-1107-8 (opens in a new tab)

  108. 108.

    Hussain G., Roychoudhury S., Singha B., Paul J. Incidence of Cryptosporidium andersoni in diarrheal patients from southern Assam, India: a molecular approach. Eur. J. Clin. Microbiol. Infect. Dis. 2017;36(6):1023-1032.

    Not in our library10.1007/s10096-016-2887-2 (opens in a new tab)

  109. 109.

    Hussein E.M., Zaki W.M., Ahmed S.A., Almatary A.M., Nemr N.I., Hussein A.M. Predominance of Giardia lamblia assemblage A among iron deficiency anaemic pre-school Egyptian children. Parasitol. Res. 2016;115:1537-1545.

    Not in our library10.1007/s00436-015-4888-y (opens in a new tab)

  110. 110.

    Ifeonu O.O., Chibucos M.C., Orvis J., Su Q., Elwin K., Guo F., Zhang H., Xiao L., Sun M., Chalmers R.M., Fraser C.M., Zhu G., Kissinger J.C., Widmer G., Silva J.C. Annotated draft genome sequences of three species of Cryptosporidium: Cryptosporidium meleagridis isolate UKMEL1, C. baileyi isolate TAMU-09Q1 and C. hominis isolates TU502_2012 and UKH1. Pathog. Dis. 2016;74.

    Not in our library10.1093/femspd/ftw080 (opens in a new tab)

  111. 111.

    Insulander M., Silverlas C., Lebbad M., Karlsson L., Mattsson J.G., Svenungsson B. Molecular epidemiology and clinical manifestations of human cryptosporidiosis in Sweden. Epidemiol. Infect. 2013;141:1009-1020.

    Not in our library10.1017/s0950268812001665 (opens in a new tab)

  112. 112.

    Iqbal J., Khalid N., Hira P.R. Cryptosporidiosis in Kuwaiti children: association of clinical characteristics with Cryptosporidium species and subtypes. J. Med. Microbiol. 2011;60:647-652.

    Not in our library10.1099/jmm.0.028001-0 (opens in a new tab)

  113. 113.

    Isaza J.P., Galvan A.L., Polanco V., Huang B., Matveyev A.V., Serrano M.G., Manque P., Buck G.A., Alzate J.F. Revisiting the reference genomes of human pathogenic Cryptosporidium species: reannotation of C. parvum Iowa and a new C. hominis reference. Sci Rep. 2015;5.

    Not in our library10.1038/srep16324 (opens in a new tab)

  114. 114.

    Jerez Puebla L.E., Nunez F.A., Martinez Silva I., Rojas Rivero L., Martinez Gonzalez M., Mendez Sutil Y., Ayllon Valdes L., Atencio Millan I., Muller N. Molecular characterization and risk factors of Giardia duodenalis among school children from La Habana, Cuba. J. Parasitol. Res. 2015;2015:378643.

    Not in our library10.1155/2015/378643 (opens in a new tab)

  115. 115.
    Genome analysis and comparative genomics of a <it>Giardia intestinalis </it>assemblage E isolate · 2010
    Not checked
  116. 116.

    Jiang J., Xiao L. An evaluation of molecular diagnostic tools for the detection and differentiation of human-pathogenic Cryptosporidium spp. J. Eukaryot. Microbiol. 2003;50:542-547.

    Not in our library10.1111/j.1550-7408.2003.tb00623.x (opens in a new tab)

  117. 117.

    Jiang J., Alderisio K.A., Xiao L. Distribution of Cryptosporidium genotypes in storm event water samples from three watersheds in New York. Appl. Environ. Microbiol. 2005;71:4446-4454.

    Not in our library10.1128/aem.71.8.4446-4454.2005 (opens in a new tab)

  118. 118.

    Jiang Y., Ren J., Yuan Z., Liu A., Zhao H., Liu H., Chu L., Pan W., Cao J., Lin Y., Shen Y. Cryptosporidium andersoni as a novel predominant Cryptosporidium species in outpatients with diarrhea in Jiangsu Province, China. BMC Infect. Dis. 2014;14:555.

    Not in our library10.1186/s12879-014-0555-7 (opens in a new tab)

  119. 119.

    Jothikumar N., da Silva A.J., Moura I., Qvarnstrom Y., Hill V.R. Detection and differentiation of Cryptosporidium hominis and Cryptosporidium parvum by dual TaqMan assays. J. Med. Microbiol. 2008;57:1099-1105.

    Not in our library10.1099/jmm.0.2008/001461-0 (opens in a new tab)

  120. 120.

    Karim M.R., Zhang S., Jian F., Li J., Zhou C., Zhang L., Sun M., Yang G., Zou F., Dong H., Li J., Rume F.I., Qi M., Wang R., Ning C., Xiao L. Multilocus typing of Cryptosporidium spp. and Giardia duodenalis from non-human primates in China. Int. J. Parasitol. 2014;44:1039-1047.

    Not in our library10.1016/j.ijpara.2014.07.006 (opens in a new tab)

  121. 121.

    Karim M.R., Wang R., Yu F., Li T., Dong H., Li D., Zhang L., Li J., Jian F., Zhang S., Rume F.I., Ning C., Xiao L. Multi-locus analysis of Giardia duodenalis from nonhuman primates kept in zoos in China: geographical segregation and host-adaptation of assemblage B isolates. Infect. Genet. Evol. 2015;30:82-88.

    Not in our library10.1016/j.meegid.2014.12.013 (opens in a new tab)

  122. 122.

    Kattula D., Jeyavelu N., Prabhakaran A.D., Premkumar P.S., Velusamy V., Venugopal S., Geetha J.C., Lazarus R.P., Das P., Nithyanandhan K., Gunasekaran C., Muliyil J., Sarkar R., Wanke C., Ajjampur S.S., Babji S., Naumova E.N., Ward H.D., Kang G. Natural history of cryptosporidiosis in a birth cohort in Southern India. Clin. Infect. Dis. 2017;64:347-354.

    Not in our library10.1093/cid/ciw730 (opens in a new tab)

  123. 123.

    Khan S.M., Debnath C., Pramanik A.K., Xiao L., Nozaki T., Ganguly S. Molecular characterization and assessment of zoonotic transmission of Cryptosporidium from dairy cattle in West Bengal, India. Vet. Parasitol. 2010;171:41-47.

    Not in our library10.1016/j.vetpar.2010.03.008 (opens in a new tab)

  124. 124.

    Koehler A.V., Whipp M.J., Haydon S.R., Gasser R.B. Cryptosporidium cuniculus - new records in human and kangaroo in Australia. Parasit. Vectors. 2014;7:492.

    Not in our library10.1186/s13071-014-0492-8 (opens in a new tab)

  125. 125.

    Korpe P.S., Haque R., Gilchrist C., Valencia C., Niu F., Lu M., Ma J.Z., Petri S.E., Reichman D., Kabir M., Duggal P., Petri W.A. Natural history of cryptosporidiosis in a longitudinal study of slum-dwelling Bangladeshi children: association with severe malnutrition. PLoS Negl. Trop. Dis. 2016;10.

    Not in our library10.1371/journal.pntd.0004564 (opens in a new tab)

  126. 126.

    Kotloff K.L., Nataro J.P., Blackwelder W.C., Nasrin D., Farag T.H., Panchalingam S., Wu Y., Sow S.O., Sur D., Breiman R.F., Faruque A.S., Zaidi A.K., Saha D., Alonso P.L., Tamboura B., Sanogo D., Onwuchekwa U., Manna B., Ramamurthy T., Kanungo S., Ochieng J.B., Omore R., Oundo J.O., Hossain A., Das S.K., Ahmed S., Qureshi S., Quadri F., Adegbola R.A., Antonio M., Hossain M.J., Akinsola A., Mandomando I., Nhampossa T., Acacio S., Biswas K., O'Reilly C.E., Mintz E.D., Berkeley L.Y., Muhsen K., Sommerfelt H., Robins-Browne R.M., Levine M.M. 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;382:209-222.

    Not in our library10.1016/s0140-6736(13)60844-2 (opens in a new tab)

  127. 127.

    Kvac M., Kvetonova D., Sak B., Ditrich O. Cryptosporidium pig genotype II in immunocompetent man. Emerg. Infect. Dis. 2009;15:982-983.

    Not in our library10.3201/eid1506.07621 (opens in a new tab)

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    Kvac M., McEvoy J., Loudova M., Stenger B., Sak B., Kvetonova D., Ditrich O., Raskova V., Moriarty E., Rost M., Macholan M., Pialek J. Coevolution of Cryptosporidium tyzzeri and the house mouse (Mus musculus). Int. J. Parasitol. 2013;43:805-817.

    Not in our library10.1016/j.ijpara.2013.04.007 (opens in a new tab)

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    Kvac M., Sakova K., Kvetonova D., Kicia M., Wesolowska M., McEvoy J., Sak B. Gastroenteritis caused by the Cryptosporidium hedgehog genotype in an immunocompetent man. J. Clin. Microbiol. 2014;52:347-349.

    Not in our library10.1128/jcm.02456-13 (opens in a new tab)

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    Learmonth J.J., Ionas G., Ebbett K.A., Kwan E.S. Genetic characterization and transmission cycles of Cryptosporidium species isolated from humans in New Zealand. Appl. Environ. Microbiol. 2004;70:3973-3978.

    Not in our library10.1128/aem.70.7.3973-3978.2004 (opens in a new tab)

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    Lebbad M., Mattsson J.G., Christensson B., Ljungstrom B., Backhans A., Andersson J.O., Svard S.G. From mouse to moose: multilocus genotyping of Giardia isolates from various animal species. Vet. Parasitol. 2010;168:231-239.

    Not in our library10.1016/j.vetpar.2009.11.003 (opens in a new tab)

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    Lebbad M., Petersson I., Karlsson L., Botero-Kleiven S., Andersson J.O., Svenungsson B., Svard S.G. 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.

    Not in our library10.1371/journal.pntd.0001262 (opens in a new tab)

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    Lebbad M., Beser J., Insulander M., Karlsson L., Mattsson J.G., Svenungsson B., Axen C. Unusual cryptosporidiosis cases in Swedish patients: extended molecular characterization of Cryptosporidium viatorum and Cryptosporidium chipmunk genotype I. Parasitology. 2013;140:1735-1740.

    Not in our library10.1017/s003118201300084x (opens in a new tab)

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    Leoni F., Amar C., Nichols G., Pedraza-Diaz S., McLauchlin J. Genetic analysis of Cryptosporidium from 2414 humans with diarrhoea in England between 1985 and 2000. J. Med. Microbiol. 2006;55:703-707.

    Not in our library10.1099/jmm.0.46251-0 (opens in a new tab)

  135. 135.
    Molecular Surveillance of Cryptosporidium spp., Giardia duodenalis, and Enterocytozoon bieneusi by Genotyping and Subtyping Parasites in Wastewater · 2012
    No notice
  136. 136.

    Li N., Xiao L., Cama V.A., Ortega Y., Gilman R.H., Guo M., Feng Y. Genetic recombination and Cryptosporidium hominis virulent subtype IbA10G2. Emerg. Infect. Dis. 2013;19:1573-1582.

    Not in our library10.3201/eid1910.121361 (opens in a new tab)

  137. 137.
    Subtyping Cryptosporidium ubiquitum, a Zoonotic Pathogen Emerging in Humans · 2014
    No notice
  138. 138.

    Li N., Neumann N.F., Ruecker N., Alderisio K.A., Sturbaum G.D., Villegas E.N., Chalmers R., Monis P., Feng Y., Xiao L. Development and evaluation of three real-time PCR assays for genotyping and source tracking Cryptosporidium spp. in water. Appl. Environ. Microbiol. 2015;81:5845-5854.

    Not in our library10.1128/aem.01699-15 (opens in a new tab)

  139. 139.
    Preliminary Characterization of MEDLE-2, a Protein Potentially Involved in the Invasion of Cryptosporidium parvum · 2017
    Not checked
  140. 140.

    Liu A., Ji H., Wang E., Liu J., Xiao L., Shen Y., Li Y., Zhang W., Ling H. Molecular identification and distribution of Cryptosporidium and Giardia duodenalis in raw urban wastewater in Harbin, China. Parasitol. Res. 2011;109:913-918.

    Not in our library10.1007/s00436-011-2333-4 (opens in a new tab)

  141. 141.
    Prevalence and genetic characterization of Cryptosporidium, Enterocytozoon, Giardia and Cyclospora in diarrheal outpatients in china · 2014
    No notice
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    Liu S., Roellig D.M., Guo Y., Li N., Frace M.A., Tang K., Zhang L., Feng Y., Xiao L. Evolution of mitosome metabolism and invasion-related proteins in Cryptosporidium. BMC Genomics. 2016;17:1006.

    Not in our library10.1186/s12864-016-3343-5 (opens in a new tab)

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    Lucio-Forster A., Griffiths J.K., Cama V.A., Xiao L., Bowman D.D. Minimal zoonotic risk of cryptosporidiosis from pet dogs and cats. Trends Parasitol. 2010;26:174-179.

    Not in our library10.1016/j.pt.2010.01.004 (opens in a new tab)

  144. 144.

    Ma J., Feng Y., Hu Y., Villegas E.N., Xiao L. Human infective potential of Cryptosporidium spp., Giardia duodenalis and Enterocytozoon bieneusi in urban wastewater treatment plant effluents. J. Water Health. 2016;14:411-423.

    Not in our library10.2166/wh.2016.192 (opens in a new tab)

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    Mallon M., MacLeod A., Wastling J., Smith H., Reilly B., Tait A. Population structures and the role of genetic exchange in the zoonotic pathogen Cryptosporidium parvum. J. Mol. Evol. 2003;56:407-417.

    Not in our library10.1007/s00239-002-2412-3 (opens in a new tab)

  146. 146.

    Mallon M.E., MacLeod A., Wastling J.M., Smith H., Tait A. Multilocus genotyping of Cryptosporidium parvum type 2: population genetics and sub-structuring. Infect. Genet. Evol. 2003;3:207-218.

    Not in our library10.1016/s1567-1348(03)00089-3 (opens in a new tab)

  147. 147.

    Mary C., Chapey E., Dutoit E., Guyot K., Hasseine L., Jeddi F., Menotti J., Paraud C., Pomares C., Rabodonirina M., Rieux A., Derouin F., Network A.C.N. Multicentric evaluation of a new real-time PCR assay for quantification of Cryptosporidium spp. and identification of Cryptosporidium parvum and Cryptosporidium hominis. J. Clin. Microbiol. 2013;51:2556-2563.

    Not in our library10.1128/jcm.03458-12 (opens in a new tab)

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    Mazurie A.J., Alves J.M., Ozaki L.S., Zhou S., Schwartz D.C., Buck G.A. Comparative genomics of Cryptosporidium. Int. J. Genomics. 2013;2013:832756.

    Not in our library10.1155/2013/832756 (opens in a new tab)

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    Mbae C., Mulinge E., Guleid F., Wainaina J., Waruru A., Njiru Z.K., Kariuki S. Molecular characterization of Giardia duodenalis in children in Kenya. BMC Infect. Dis. 2016;16:135.

    Not in our library10.1186/s12879-016-1436-z (opens in a new tab)

  150. 150.

    Minetti C., Lamden K., Durband C., Cheesbrough J., Fox A., Wastling J.M. Determination of Giardia duodenalis assemblages and multi-locus genotypes in patients with sporadic giardiasis from England. Parasit. Vectors. 2015;8:444.

    Not in our library10.1186/s13071-015-1059-z (opens in a new tab)

  151. 151.

    Minetti C., Lamden K., Durband C., Cheesbrough J., Platt K., Charlett A., O'Brien S.J., Fox A., Wastling J.M. Case-control study of risk factors for sporadic giardiasis and parasite assemblages in North West England. J. Clin. Microbiol. 2015;53:3133-3140.

    Not in our library10.1128/jcm.00715-15 (opens in a new tab)

  152. 152.

    Mohammed Mahdy A.K., Surin J., Wan K.L., Mohd-Adnan A., Al-Mekhlafi M.S., Lim Y.A. Giardia intestinalis genotypes: risk factors and correlation with clinical symptoms. Acta Trop. 2009;112:67-70.

    Not in our library10.1016/j.actatropica.2009.06.012 (opens in a new tab)

  153. 153.

    Molloy S.F., Smith H.V., Kirwan P., Nichols R.A., Asaolu S.O., Connelly L., Holland C.V. Identification of a high diversity of Cryptosporidium species genotypes and subtypes in a pediatric population in Nigeria. Am. J. Trop. Med. Hyg. 2010;82:608-613.

    Not in our library10.4269/ajtmh.2010.09-0624 (opens in a new tab)

  154. 154.

    Monis P.T., Caccio S.M., Thompson R.C. Variation in Giardia: towards a taxonomic revision of the genus. Trends Parasitol. 2009;25:93-100.

    Not in our library10.1016/j.pt.2008.11.006 (opens in a new tab)

  155. 155.

    Moore C.E., Elwin K., Phot N., Seng C., Mao S., Suy K., Kumar V., Nader J., Bousfield R., Perera S., Bailey J.W., Beeching N.J., Day N.P., Parry C.M., Chalmers R.M. Molecular characterization of Cryptosporidium species and Giardia duodenalis from symptomatic Cambodian children. PLoS Negl. Trop. Dis. 2016;10.

    Not in our library10.1371/journal.pntd.0004822 (opens in a new tab)

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    Moreira N.A., Bondelind M. Safe drinking water and waterborne outbreaks. J. Water Health. 2017;15:83-96.

    Not in our library10.2166/wh.2016.103 (opens in a new tab)

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    Morrison H.G., McArthur A.G., Gillin F.D., Aley S.B., Adam R.D., Olsen G.J., Best A.A., Cande W.Z., Chen F., Cipriano M.J., Davids B.J., Dawson S.C., Elmendorf H.G., Hehl A.B., Holder M.E., Huse S.M., Kim U.U., Lasek-Nesselquist E., Manning G., Nigam A., Nixon J.E., Palm D., Passamaneck N.E., Prabhu A., Reich C.I., Reiner D.S., Samuelson J., Svard S.G., Sogin M.L. Genomic minimalism in the early diverging intestinal parasite Giardia lamblia. Science. 2007;317:1921-1926.

    Not in our library10.1126/science.1143837 (opens in a new tab)

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    Morrison L.J., Mallon M.E., Smith H.V., Macleod A., Xiao L., Tait A. The population structure of the Cryptosporidium parvum population in Scotland: a complex picture. Infect. Genet. Evol. 2008;8:121-129.

    Not in our library10.1016/j.meegid.2007.10.010 (opens in a new tab)

  159. 159.

    Morse T.D., Nichols R.A., Grimason A.M., Campbell B.M., Tembo K.C., Smith H.V. Incidence of cryptosporidiosis species in paediatric patients in Malawi. Epidemiol. Infect. 2007;135:1307-1315.

    Not in our library10.1017/s0950268806007758 (opens in a new tab)

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    Nazemalhosseini-Mojarad E., Feng Y., Xiao L. The importance of subtype analysis of Cryptosporidium spp. in epidemiological investigations of human cryptosporidiosis in Iran and other Mideast countries. Gastroenterol. Hepatol. Bed Bench. 2012;5:67-70.

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    Neira O.P., Munoz S.N., Wilson L.G., Barthel M.M., Rosales L.M., Henriquez R.C. Cryptosporidium species in immunodeficient and immunocompetent patients of Valparaiso: a descriptive study. Rev. Chil. Infectol. 2012;29:63-71.

    Not in our library10.4067/s0716-10182012000100011 (opens in a new tab)

  162. 162.

    Network A.C.N. Laboratory-based surveillance for Cryptosporidium in France, 2006–2009. Euro Surveill. 2010;15.

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  163. 163.

    Ng J., MacKenzie B., Ryan U. Longitudinal multi-locus molecular characterisation of sporadic Australian human clinical cases of cryptosporidiosis from 2005 to 2008. Exp. Parasitol. 2010;125:348-356.

    Not in our library10.1016/j.exppara.2010.02.017 (opens in a new tab)

  164. 164.

    Ng J.S., Eastwood K., Walker B., Durrheim D.N., Massey P.D., Porigneaux P., Kemp R., McKinnon B., Laurie K., Miller D., Bramley E., Ryan U. Evidence of Cryptosporidium transmission between cattle and humans in northern New South Wales. Exp. Parasitol. 2012;130:437-441.

    Not in our library10.1016/j.exppara.2012.01.014 (opens in a new tab)

  165. 165.

    Ng-Hublin J.S., Combs B., Mackenzie B., Ryan U. Human cryptosporidiosis diagnosed in Western Australia – a mixed infection with Cryptosporidium meleagridis, Cryptosporidium mink genotype and an unknown Cryptosporidium species. J. Clin. Microbiol. 2013;51:2463-2465.

    Not in our library10.1128/jcm.00424-13 (opens in a new tab)

  166. 166.

    Ng-Hublin J.S.Y., Combs B., Reid S., Ryan U. Differences in the occurrence and epidemiology of cryptosporidiosis in Aboriginal and non-Aboriginal people in Western Australia (2002–2012). Infect. Genet. Evol. 2017;53:100-106.

    Not in our library10.1016/j.meegid.2017.05.018 (opens in a new tab)

  167. 167.

    Operario D.J., Bristol L.S., Liotta J., Nydam D.V., Houpt E.R. Correlation between diarrhea severity and oocyst count via quantitative PCR or fluorescence microscopy in experimental cryptosporidiosis in calves. Am. J. Trop. Med. Hyg. 2015;92:45-49.

    Not in our library10.4269/ajtmh.14-0488 (opens in a new tab)

  168. 168.

    Painter J.E., Hlavsa M.C., Collier S.A., Xiao L., Yoder J.S. Cryptosporidiosis surveillance - United States, 2011–2012. MMWR Surveill. Summ. 2015;64(Suppl. 3):1-14.

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    Painter J.E., Gargano J.W., Yoder J.S., Collier S.A., Hlavsa M.C. Evolving epidemiology of reported cryptosporidiosis cases in the United States, 1995–2012. Epidemiol. Infect. 2016;144:1792-1802.

    Not in our library10.1017/s0950268815003131 (opens in a new tab)

  170. 170.

    Palmer C.J., Xiao L., Terashima A., Guerra H., Gotuzzo E., Saldias G., Bonilla J.A., Zhou L., Lindquist A., Upton S.J. Cryptosporidium muris, a rodent pathogen, recovered from a human in Peru. Emerg. Infect. Dis. 2003;9:1174-1176.

    Not in our library10.3201/eid0909.030047 (opens in a new tab)

  171. 171.

    Pantchev N., Broglia A., Paoletti B., Globokar Vrhovec M., Bertram A., Nockler K., Caccio S.M. Occurrence and molecular typing of Giardia isolates in pet rabbits, chinchillas, guinea pigs and ferrets collected in Europe during 2006–2012. Vet. Rec. 2014;175:18.

    Not in our library10.1136/vr.102236 (opens in a new tab)

  172. 172.

    Paparini A., Gofton A., Yang R., White N., Bunce M., Ryan U.M. Comparison of Sanger and next generation sequencing performance for genotyping Cryptosporidium isolates at the 18S rRNA and actin loci. Exp. Parasitol. 2015;151–152:21-27.

    Not in our library10.1016/j.exppara.2015.02.001 (opens in a new tab)

  173. 173.

    Pelayo L., Nunez F.A., Rojas L., Furuseth Hansen E., Gjerde B., Wilke H., Mulder B., Robertson L. Giardia infections in Cuban children: the genotypes circulating in a rural population. Ann. Trop. Med. Parasitol. 2008;102:585-595.

    Not in our library10.1179/136485908x355247 (opens in a new tab)

  174. 174.

    Perez Cordon G., Cordova Paz Soldan O., Vargas Vasquez F., Velasco Soto J.R., Sempere Bordes L., Sanchez Moreno M., Rosales M.J. Prevalence of enteroparasites and genotyping of Giardia lamblia in Peruvian children. Parasitol. Res. 2008;103:459-465.

    Not in our library10.1007/s00436-008-1007-3 (opens in a new tab)

  175. 175.

    Perez-Cordon G., Robinson G., Nader J., Chalmers R.M. Discovery of new variable number tandem repeat loci in multiple Cryptosporidium parvum genomes for the surveillance and investigation of outbreaks of cryptosporidiosis. Exp. Parasitol. 2016;169:119-128.

    Not in our library10.1016/j.exppara.2016.08.003 (opens in a new tab)

  176. 176.

    Petrincova A., Valencakova A., Luptakova L., Ondriska F., Kalinova J., Halanova M., Danisova O., Jarcuska P. Molecular characterization and first report of Cryptosporidium genotypes in human population in the Slovak Republic. Electrophoresis. 2015;36:1023-1032.

    Not in our library10.1002/elps.201500230 (opens in a new tab)

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    Pijnacker R., Mughini-Gras L., Heusinkveld M., Roelfsema J., van Pelt W., Kortbeek T. Different risk factors for infection with Giardia lamblia assemblages A and B in children attending day-care centres. Eur. J. Clin. Microbiol. Infect. Dis. 2016;35:2005-2013.

    Not in our library10.1007/s10096-016-2753-2 (opens in a new tab)

  178. 178.
    Pathogen-specific burdens of community diarrhoea in developing countries: a multisite birth cohort study (MAL-ED) · 2015
    Not checked
  179. 179.

    Prystajecky N., Tsui C.K., Hsiao W.W., Uyaguari-Diaz M.I., Ho J., Tang P., Isaac-Renton J. Giardia spp. are commonly found in mixed assemblages in surface water, as revealed by molecular and whole-genome characterization. Appl. Environ. Microbiol. 2015;81:4827-4834.

    Not in our library10.1128/aem.00524-15 (opens in a new tab)

  180. 180.

    Puleston R.L., Mallaghan C.M., Modha D.E., Hunter P.R., Nguyen-Van-Tam J.S., Regan C.M., Nichols G.L., Chalmers R.M. The first recorded outbreak of cryptosporidiosis due to Cryptosporidium cuniculus (formerly rabbit genotype), following a water quality incident. J. Water Health. 2014;12:41-50.

    Not in our library10.2166/wh.2013.097 (opens in a new tab)

  181. 181.

    Qi M., Xi J., Li J., Wang H., Ning C., Zhang L. Prevalence of zoonotic Giardia duodenalis assemblage B and first identification of assemblage E in rabbit fecal samples isolates from Central China. J. Eukaryot. Microbiol. 2015;62:810-814.

    Not in our library10.1111/jeu.12239 (opens in a new tab)

  182. 182.

    Qi M., Yu F., Li S., Wang H., Luo N., Huang J., Zhang L. Multilocus genotyping of potentially zoonotic Giardia duodenalis in pet chinchillas (Chinchilla lanigera) in China. Vet. Parasitol. 2015;208:113-117.

    Not in our library10.1016/j.vetpar.2015.02.001 (opens in a new tab)

  183. 183.

    Qi M., Wang H., Jing B., Wang R., Jian F., Ning C., Zhang L. Prevalence and multilocus genotyping of Giardia duodenalis in dairy calves in Xinjiang, Northwestern China. Parasit. Vectors. 2016;9:546.

    Not in our library10.1186/s13071-016-1828-3 (opens in a new tab)

  184. 184.

    Qi M., Wang R., Jing B., Jian F., Ning C., Zhang L. Prevalence and multilocus genotyping of Cryptosporidium andersoni in dairy cattle and He cattle in Xinjiang, China. Infect. Genet. Evol. 2016;44:313-317.

    Not in our library10.1016/j.meegid.2016.07.022 (opens in a new tab)

  185. 185.

    Quilez J., Vergara-Castiblanco C., Monteagudo L., Del Cacho E., Sanchez-Acedo C. Multilocus fragment typing and genetic structure of Cryptosporidium parvum isolates from diarrheic preweaned calves in Spain. Appl. Environ. Microbiol. 2011;77:7779-7786.

    Not in our library10.1128/aem.00751-11 (opens in a new tab)

  186. 186.

    Quilez J., Vergara-Castiblanco C., Monteagudo L., del Cacho E., Sanchez-Acedo C. Host association of Cryptosporidium parvum populations infecting domestic ruminants in Spain. Appl. Environ. Microbiol. 2013;79:5363-5371.

    Not in our library10.1128/aem.01168-13 (opens in a new tab)

  187. 187.

    Ramo A., Monteagudo L.V., Del Cacho E., Sanchez-Acedo C., Quilez J. Intra-species genetic diversity and clonal structure of Cryptosporidium parvum in sheep farms in a confined geographical area in Northeastern Spain. PLoS ONE. 2016;11.

    Not in our library10.1371/journal.pone.0155336 (opens in a new tab)

  188. 188.

    Ramo A., Quilez J., Monteagudo L., Del Cacho E., Sanchez-Acedo C. Intra-species diversity and panmictic structure of Cryptosporidium parvum populations in cattle farms in Northern Spain. PLoS ONE. 2016;11.

    Not in our library10.1371/journal.pone.0148811 (opens in a new tab)

  189. 189.

    Ramo A., Del Cacho E., Sanchez-Acedo C., Quilez J. Occurrence and genetic diversity of Cryptosporidium and Giardia in urban wastewater treatment plants in north-eastern Spain. Sci. Total Environ. 2017;598:628-638.

    Not in our library10.1016/j.scitotenv.2017.04.097 (opens in a new tab)

  190. 190.

    Raskova V., Kvetonova D., Sak B., McEvoy J., Edwinson A., Stenger B., Kvac M. Human cryptosporidiosis caused by Cryptosporidium tyzzeri and C. parvum isolates presumably transmitted from wild mice. J. Clin. Microbiol. 2013;51:360-362.

    Not in our library10.1128/jcm.02346-12 (opens in a new tab)

  191. 191.

    Read C., Walters J., Robertson I.D., Thompson R.C. Correlation between genotype of Giardia duodenalis and diarrhoea. Int. J. Parasitol. 2002;32:229-231.

    Not in our library10.1016/s0020-7519(01)00340-x (opens in a new tab)

  192. 192.

    Robinson G., Elwin K., Chalmers R.M. Unusual Cryptosporidium genotypes in human cases of diarrhea. Emerg. Infect. Dis. 2008;14:1800-1802.

    Not in our library10.3201/eid1411.080239 (opens in a new tab)

  193. 193.

    Roelfsema J.H., Sprong H., Caccio S.M., Takumi K., Kroes M., van Pelt W., Kortbeek L.M., van der Giessen J.W. Molecular characterization of human Cryptosporidium spp. isolates after an unusual increase in late summer 2012. Parasit. Vectors. 2016;9:138.

    Not in our library10.1186/s13071-016-1397-5 (opens in a new tab)

  194. 194.

    Ryan U., Caccio S.M. Zoonotic potential of Giardia. Int. J. Parasitol. 2013;43:943-956.

    Not in our library10.1016/j.ijpara.2013.06.001 (opens in a new tab)

  195. 195.

    Ryan U., Xiao L., Read C., Zhou L., Lal A.A., Pavlasek I. Identification of novel Cryptosporidium genotypes from the Czech Republic. Appl. Environ. Microbiol. 2003;69:4302-4307.

    Not in our library10.1128/aem.69.7.4302-4307.2003 (opens in a new tab)

  196. 196.

    Ryan U., Fayer R., Xiao L. Cryptosporidium species in humans and animals: current understanding and research needs. Parasitology. 2014;141:1667-1685.

    Not in our library10.1017/s0031182014001085 (opens in a new tab)

  197. 197.

    Ryan U., Paparini A., Oskam C. New technologies for detection of enteric parasites. Trends Parasitol. 2017;33(7):532-546.

    Not in our library10.1016/j.pt.2017.03.005 (opens in a new tab)

  198. 198.

    Sahagun J., Clavel A., Goni P., Seral C., Llorente M.T., Castillo F.J., Capilla S., Arias A., Gomez-Lus R. Correlation between the presence of symptoms and the Giardia duodenalis genotype. Eur. J. Clin. Microbiol. Infect. Dis. 2008;27:81-83.

    Not in our library10.1007/s10096-007-0404-3 (opens in a new tab)

  199. 199.

    Samie A., Ntekele P. Genotypic detection and evaluation of the removal efficiency of Giardia duodenalis at municipal wastewater treatment plants in Northern South Africa. Trop. Biomed. 2014;31:122-133.

    Not in our library

  200. 200.

    Sanchez A., Munoz M., Gomez N., Tabares J., Segura L., Salazar A., Restrepo C., Ruiz M., Reyes P., Qian Y., Xiao L., Lopez M.C., Ramirez J.D. Molecular epidemiology of Giardia, Blastocystis and Cryptosporidium among indigenous children from the Colombian Amazon basin. Front. Microbiol. 2017;8:248.

    Not in our library10.3389/fmicb.2017.00248 (opens in a new tab)

  201. 201.

    Santin M., Cortes Vecino J.A., Fayer R. A large scale molecular study of Giardia duodenalis in horses from Colombia. Vet. Parasitol. 2013;196:31-36.

    Not in our library10.1016/j.vetpar.2013.02.006 (opens in a new tab)

  202. 202.

    Scalia L.A., Fava N.M., Soares R.M., Limongi J.E., da Cunha M.J., Pena I.F., Kalapothakis E., Cury M.C. Multilocus genotyping of Giardia duodenalis in Brazilian children. Trans. R. Soc. Trop. Med. Hyg. 2016;110:343-349.

    Not in our library10.1093/trstmh/trw036 (opens in a new tab)

  203. 203.

    Sikora P., Andersson S., Winiecka-Krusnell J., Hallstrom B., Alsmark C., Troell K., Beser J., Arrighi R.B. Genomic variation in IbA10G2 and other patient-derived Cryptosporidium hominis subtypes. J. Clin. Microbiol. 2017;55:844-858.

    Not in our library10.1128/jcm.01798-16 (opens in a new tab)

  204. 204.

    Skhal D., Aboualchamat G., Al Mariri A., Al Nahhas S. Prevalence of Giardia duodenalis assemblages and sub-assemblages in symptomatic patients from Damascus city and its suburbs. Infect. Genet. Evol. 2017;47:155-160.

    Not in our library10.1016/j.meegid.2016.11.030 (opens in a new tab)

  205. 205.

    Soliman R.H., Fuentes I., Rubio J.M. Identification of a novel assemblage B subgenotype and a zoonotic assemblage C in human isolates of Giardia intestinalis in Egypt. Parasitol. Int. 2011;60:507-511.

    Not in our library10.1016/j.parint.2011.09.006 (opens in a new tab)

  206. 206.
    Identification of Zoonotic Genotypes of Giardia duodenalis · 2009
    No notice
  207. 207.
    Cryptosporidium and Giardia in Africa: current and future challenges · 2017
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