Standardization of molecular techniques for the detection and characterization of intestinal protozoa and other pathogens in humans

Maria Alejandra Vethencourt Ysea Mariana Cedeño Umaña Sofia Pereira Fuentes Idalia Valerio Campos Misael Chinchilla Carmona About the authors

Abstract

Background:

The intrinsic sensitivity limitations of basic parasitological methods, along with the particular biological characteristics of parasites, make these methods ineffective to differentiate morphologically indistinguishable species. Molecular detection and characterization techniques could be used to overcome these problems. The purpose of this work was to standardize molecular polymerase chain reaction (PCR) techniques, described in the literature, for the detection and molecular characterization of intestinal protozoa and other pathogens in humans.

Methods:

DNA was extracted from human or animal feces, previously washed or cultured in Boeck Drbohlav's Modified Medium. DNA extraction was performed with Machery-Nagel extraction kits. The standardization of the PCR, nested-PCR or RFLP techniques was carried out according to the literature. For each molecular technique performed, the sensitivity of the test was determined based on the minimun quantity required of DNA (sensitivity A) and the minimum quantity of life forms that the test detected (sensitivity B).

Results:

Sensitivity A was 10 fg for G. duodenalis, 12.5 pg for Entamoeba histolytica or Entamoeba dispar, 50 fg for Cryptosporidium spp., 225 pg for Cyclospora spp. and 800 fg or 8 fg for Blastocystis spp. after performing a 1780 bp PCR or 310 bp nested PCR, respectively. The sensitivity B was 100 cysts for G. duodenalis, 500 cysts for E. histolytica or E. dispar, 1000 oocysts for Cyclospora spp. and 3600 or four vegetatives forms for PCR or nested PCR of Blastocystis spp., respectively.

Conclusions:

The molecular detection of protozoa and chromist was achieved and the molecular characterization allowed the genotyping of some of the parasites such as Giardia duodenalis, Cryptosporidium spp., and Blastocystis spp. This study summarizes the molecular techniques for epidemiological studies in humans and animals, and helps in the investigation of their transmission sources in countries where intestinal parasites are a public health problem.

Keywords:
Molecular detection; Molecular characterization; PCR-RFLP; Intestinal protozoa; Blastocystis spp.; Costa Rica

Background

Intestinal human parasites have a worldwide distribution. They can be protozoa and helminths and are a public health problem that affects people of all ages, especially children and the elderly, particularly in least developed countries [11. Chelkeba L, Mekonnen Z, Alemu Y, Emana D. Epidemiology of intestinal parasitic infections in preschool and school-aged Ethiopian children: a systematic review and meta-analysis. BMC Public Health. 2020 Jan 28;20(1):117.]. According to the 2008-2009 National Health Survey [22. Costa Rica. Ministerio de Salud. Instituto Costarricense de Investigación y Enseñanza en Nutrición y Salud. Caja Costarricense de Seguro Social. Instituto Nacional de Estadística y Censos. Instituto Costarricense Sobre Drogas. Encuesta Nacional de Nutrición 2008-2009. Fascículo 5: Parásitos Intestinales. San José, Costa Rica: El Ministerio, 2012. Available from: https://www.binasss.sa.cr/opac-ms/media/digitales/Encuesta%20Nacional%20de%20Nutrici%C3%B3n.%20Fasc%C3%ADculo%205.%20Par%C3%A1sitos%20intestinales.pdf.
https://www.binasss.sa.cr/opac-ms/media/...
], the prevalence of Blastocystis spp. in Costa Rica was around 37.8%, increasing with age to 46.6%. Entamoeba histolytica/ E. dispar leads the list among the pathogenic protozoans with 3.2% followed by Giardia duodenalis with 2.9% already. The eradication of intestinal helminths in Costa Rica was recorded [33. Mata L. Erradicación de helmintos intestinales en Costa Rica. Rev Med Hosp Nac Niños Costa Rica. 1985;20(2):151-64. ]; however, in more recent studies carried out by Abrahams-Sandi et al. [44. Abrahams-Sandí E, Solano M., Rodríguez B. Prevalencia de parásitos intestinales en escolares de Limón Centro, Costa Rica. Rev Costarric Cienc Med. 2005;26(1-2):33-8. ] and Arévalo et al. [55. Arévalo M, Cortés X, Barrantes K, Achí R. Prevalence of intestinal parasitosis in children from the community of Los Cuadros, Goicoechea, Costa Rica. 2002-2003. Rev Costarric Cienc Med. 2007 Jun;28(1-2):37-45.], higher prevalences of helminths and of pathogenic protozoa were present in children in Limón [44. Abrahams-Sandí E, Solano M., Rodríguez B. Prevalencia de parásitos intestinales en escolares de Limón Centro, Costa Rica. Rev Costarric Cienc Med. 2005;26(1-2):33-8. ] and Goicochea [55. Arévalo M, Cortés X, Barrantes K, Achí R. Prevalence of intestinal parasitosis in children from the community of Los Cuadros, Goicoechea, Costa Rica. 2002-2003. Rev Costarric Cienc Med. 2007 Jun;28(1-2):37-45.]. According to the 2008-2009 National Survey [22. Costa Rica. Ministerio de Salud. Instituto Costarricense de Investigación y Enseñanza en Nutrición y Salud. Caja Costarricense de Seguro Social. Instituto Nacional de Estadística y Censos. Instituto Costarricense Sobre Drogas. Encuesta Nacional de Nutrición 2008-2009. Fascículo 5: Parásitos Intestinales. San José, Costa Rica: El Ministerio, 2012. Available from: https://www.binasss.sa.cr/opac-ms/media/digitales/Encuesta%20Nacional%20de%20Nutrici%C3%B3n.%20Fasc%C3%ADculo%205.%20Par%C3%A1sitos%20intestinales.pdf.
https://www.binasss.sa.cr/opac-ms/media/...
], the prevalence of parasites in Costa Rica is closely related to poor socio-economic and hygienic characteristics [66. Solano-Barquero M, Montero-Salguero A, León-Alán D, Santamaría-Ulloa C, Mora A M, Reyes-Lizano L. Prevalence of parasitosis in children aged 1 to 7 years in a vulnerable condition in the Central-South Region of Costa Rica. Acta Med Costarric. 2018 Apr-Jun;60(2):19-29.].

Parasitic bowel diseases cause low mortality; however, the easy transmission way, the chronicity of symptoms, and the associated potential complications are important public health and sanitary problems. Some intestinal parasites can go unnoticed without producing symptoms, but they can also cause digestive symptoms of varying intensity, even with serious repercussions on the body [77. Solano L, Acuña I, Barón M, Morón DE SA, Sánchez, A. Intestinal parasitic infestation and others infectious backgrounds influence in the antropometric nutritional status of children in poverty. Parasitol Latinoam. 2008;63(1-4):12-9. ]. They can affect the individual’s productivity or educational performance, causing absenteeism from work, and anthropometric nutritional status in infected school-age children [77. Solano L, Acuña I, Barón M, Morón DE SA, Sánchez, A. Intestinal parasitic infestation and others infectious backgrounds influence in the antropometric nutritional status of children in poverty. Parasitol Latinoam. 2008;63(1-4):12-9. ].

Detection techniques that are based on basic parasitological methods such as direct examination, concentration or culture methods have sensitivity limitations. Moreover, the particular biological characteristics of the parasites make them more inefficient when it comes to detecting low counts or to differentiate species that are morphologically indistinguishable [88. Akgül Ö, Kart Yaşar K, Sapmaz B, Kırkoyun Uysal H, Yıldırmak T, Şimşek F, Karasakal ÖF, Çalışkan R, Öner YA. [Detection of intestinal parasites with conventional and molecular methods in follow-up HIV/AIDS cases]. Mikrobiyol Bul. 2018 Jul;52(3):273-83. [Article in Turkish].,99. Jerez Puebla LE, Núñez-Fernández FA, Fraga Nodarse J, Atencio Millán I, Cruz Rodríguez I, Martínez Silva I, Ayllón Valdés L, Robertson LJ. Diagnosis of intestinal protozoan infections in patients in Cuba by microscopy and molecular methods: advantages and disadvantages. J Microbiol Methods. 2020 Dec;179:106102.]. Cases like these require the sensitivity characteristics of molecular biology techniques.

The application of molecular techniques to detect and identify the DNA of the parasite sets the scene for the molecular epidemiology of intestinal parasitosis, especially for pathogens that are morphologically indistinguishable from commensal species. An example of this would be the differentiation of Entamoeba histolytica (pathogen) from the commensal species (Entamoeba dispar or Entamoeba moshkovskii) morphologically indistinguishable [1010. Rivero de Rodríguez Z. Detection of Entamoeba moshkovskii in humans: a new diagnostic problem in amebiasis diagnosis. Kasmera. 2013;41(1):42-9.]. In the case of the genus Blastocystis spp., genotypes are morphologically indistinguishable, but some of them have been related to pathogenicity [1111. Salehi R, Haghighi A, Stensvold CR, Kheirandish F, Azargashb E, Raeghi S, Kohansal C, Bahrami F. Prevalence and subtype identification of Blastocystis isolated from humans in Ahvaz, Southwestern Iran. Gastroenterol Hepatol Bed Bench. 2017;10(3):235-41.]. Molecular biology allows the detection Cryptosporidium spp., a small size protozoan that requires special staining, such as the modified Ziehl Neelsen [1212. Tahvildar-Biderouni F, Salehi N. Detection of Cryptosporidium infection by modified ziehl-neelsen and PCR methods in children with diarrheal samples in pediatric hospitals in Tehran. Gastroenterol Hepatol Bed Bench. 2014;7(2):125-30.] technique, to observe its presence. The use of molecular tools would increase the detection level to make it possible to identify the presence of these species. Currently, 42 species of Cryptosporidium are recognized [1313. Zahedi A, Ryan U. Cryptosporidium - An update with an emphasis on foodborne and waterborne transmission. Res Vet Sci. 2020 Oct;132:500-12.], with Cryptosporidium parvum and Cryptosporidium hominis being responsible for more than 90% of human infections [1414. Feng Y, Ryan UM, Xiao L. Genetic Diversity and Population Structure of Cryptosporidium. Trends Parasitol. 2018 Nov;34(11):997-1011.]. C. hominis has been linked to anthroponotic tranmission [1515. Xiao L, Feng Y. Molecular epidemiologic tools for waterborne pathogens Cryptosporidium spp. and Giardia duodenalis. Food Waterborne Parasitol. 2017 Sep 29;8-9:14-32.] whereas C. parvum presents a zoonotic transmission route with livestock as the primary source of infection [1616. Nader JL, Mathers TC, Ward BJ, Pachebat JA, Swain MT, Robinson G, Chalmers RM, Hunter PR, van Oosterhout C, Tyler KM. Evolutionary genomics of anthroponosis in Cryptosporidium. Nat Microbiol. 2019 May;4(5):826-36.]. Therefore, the application of molecular techniques would allow the identification of human pathogenic intestinal protozoa, increase the detection limit, and even improve the characterization of species or genotypes.

In this work, a compendium of molecular biology techniques that allow the detection and molecular characterization of the genus and species or genotypes of protozoa pathogens in humans is presented. Moreover, the detection limit of evolutionary life forms of E. histolytica and/or E. dispar and/or E. moshkovskii, G. duodenalis, Cryptosporidium spp., Cyclospora spp. and stramenopiles such as Blastocystis spp. is also suggested as a molecular detection tool for parasites.

The present study paves the way for the molecular epidemiology of intestinal protozoosis and Blastocystis spp., which may serve for epidemiological studies in humans and animals, the investigation of the sources of transmission of some of these parasites and as diagnostic tools, which may be used at the service of the population.

Methods

Obtaining positive controls for molecular tests

The parasites to be detected molecularly were obtained from human and animal feces, donated to UCIMED Basic Research Laboratory (LIB-UCIMED). Direct parasitological analysis was performed on the samples with 0.85% saline and lugol [1717. Guzmán de Rondón C, Vethencourt MA, Galindo M, Chacón N, Wagner C, Nessi A. Comportamiento biológico de Blastocystis hominis en pacientes tratados con Secnidazol (Unidazol®) / Biological behavior of Blastocystis hominis in patients treated with Secnidazol (Unidazol®). Rev Soc Venez Microbiol. 2008 Jun;28(1):66-71. ] looking for protozoan cysts or with permanent stains such as modified Koster to detect Cryptosporidium spp. [1818. Kageruka P, Brandt JR, Taelman H, Jonas C. Modified Köster staining method for the diagnosis of cryptosporidiosis. Ann Soc Belg Med Trop. 1984 Jun;64(2):171-5.], or Ziehl Nielsen to observe the presence of Cyclospora spp. [1919. Murray PR, Baron EJ. Manual of Clinical Microbiology. 9th ed. Murray PR, editor. Washington, D.C: ASM Press; 2007.]. The feces samples with cysts, were washed (HL) to eliminate the excess of contaminants inherent in the sample, resuspending 1 g of the sample in 10 mL sterile warm distilled water, macerating with wooden sticks and centrifuged at 1750 x g/10 min (Frontier™ Serie 5000, OHAUS, USA). The procedure was carried out as many times as necessary to obtain a clear supernatant. The final pellet was resuspended in 5 mL of sterile distilled water, aliquoted at a rate of 1 mL in 1.5 mL ependorf tubes, centrifuged at 21380 x g/2 min (Mikro 200, Hettich Instruments) and stored at -20 °C until DNA extraction. DNA extraction was performed with the Machery-Nagel Extraction Kit (NucleoSpin® Tissue). Feces samples positive for Blastocystis spp. (vegetative forms and/or cysts) were cultivated in Modified Boeck Drbohlav's Medium (MBDM) for up to 96 hours to increase the vegetative forms of this parasite [1717. Guzmán de Rondón C, Vethencourt MA, Galindo M, Chacón N, Wagner C, Nessi A. Comportamiento biológico de Blastocystis hominis en pacientes tratados con Secnidazol (Unidazol®) / Biological behavior of Blastocystis hominis in patients treated with Secnidazol (Unidazol®). Rev Soc Venez Microbiol. 2008 Jun;28(1):66-71. ]. Finally, the pellet was washed 3 times with Ringer's buffer by centrifugation at 1750 × g/10 min, and subreadsequently stored at -20° C until use.

DNA extraction

The DNA extraction was carried out with the Machery-Nagel extraction kits (NucleoSpin® Tissue), standardizing the mechanical lysis process with and without the presence of glass beads, according to the suggestions of Sepahvand et al. [2020. Sepahvand A, Pestehchian N, Yousefi HA, Gharehbaba RP. Comparison and evaluation of four methods for extracting DNA from Giardia duodenalis cysts for PCR targeting the tpi gene. J Parasit Dis. 2017 Mar;41(1):263-7.]. Briefly, the pellet stored at -20° C was resuspended with 250 µL of TE buffer (10 mM Tris/HCl; 1 mM EDTA, pH 8) plus 200 mg of cover glass powder #1, sterile. The lysis process was carried out three times. Each cycle consisted of cooling the preparation for 3 min at 4° C in a thermal block (Torrey Pines Scientic, USA) and mixing for 3 min in vortex. Then, it was centrifuged for 1 min at 21380 x g/2 min (Mikro 200, Hettich Instruments), the supernatant was transferred to a clean tube, and the DNA extraction was continued with the Machery-Nagel Kits (NucleoSpin® Tissue), following the indications of the commercial company recommended for the extraction of eukaryotic cells.

Measurement of DNA quality

DNA was quantified with the Qubit fluorometer (ThermoFisher, USA), following the manufacturer's instructions. The integrity of the DNA (10 µL), with 2 µL of the 6X loading buffer (6X MassRuler, Loading Dye Solution, Fermentas) was evaluated after carrying out an agarose gel electrophoresis (SeaKem LE, Cambrex, USA) at 1%, dissolved in Tris-Acetate-EDTA Buffer (TAE), stained with GelRed (Gel Stain, Biotium, Cat: 41003). Electrophoresis was performed at 100 volts (FB1000 Power Source, Fisher Scientific, USA). To visualize the PCR amplifications and the RFLPs, electrophoresis was carried out on a 2% agarose gel, following the same methodology. The size of the amplified PCR, and of its fragments obtained after digestion with the enzymes (RFLP), was compared with a marker of 50 bp (DNA ladder GeneRuler, #SM0371, ThermoFisher, USA) or 100 bp (DNA ladder GeneRuler, #SM0241, ThermoFisher, USA). Image analysis was performed with a UV transilluminator (Slimline Series; Spectroline), the image was captured with an image digitizer (Enduro™ GDS, Labnet International, Inc., USA). Both the confirmation of the size of the amplified by PCR and the analysis of the RFLP were carried out with the TotalLab 1D software, version 14.0.

Molecular techniques for the detection of pathogens

The PCR, nested-PCR or RFLP techniques were carried out according to the suggestion of the references for each of the parasites to be studied. For each molecular test the optimal concentration of primers, deoxyribonucleotide triphosphates (Dntps; Thermo Scientific, Cat. R0191, USA), Magnesium chloride (MgCl2; Thermo Scientific, Lot. 00603943, USA) and DNA polymerase (DreamTaq, DNA polymerase; Thermo Scientific, Cat. EP0702, USA) were adjusted and standardized to achieve a single amplification of the size suggested by the literature. For each of the molecular techniques performed, the sensitivity of the test was determined based on the minimun quantity required of DNA (sensitibity A) and the fewest number of life forms that the test detected (sensitibity B). Sensitivity A was performed from factor 10 dilutions of a DNA sample of known concentration and extracted from a feces sample whose positivity was verified by parasitological examination. Sensitivity B was performed from serial dilutions, by factor 10, of a fecal sample with a known count of vegetative forms, cysts or oocysts, followed by DNA extraction. The digestions with the restriction enzymes were performed at a final volume of 25 μL, adjusted with nuclease-free water, BE 1X buffer (Buffer CutSmart, BioLabs, New England, USA). Simple or double digestion was performed with 10 units of each of the fast-digesting restriction enzymes (BioLabs, New England, USA) and 5 to 10 μL of the amplification obtained by PCR at 37 °C/15 min. Table 1 summarizes the type of molecular technique used, the gene to amplify, the species to be defined, the name of the primers and their sequence, the size of the amplified, the type of restriction enzymes used, and the base references of each one of the standardized tests. Table 2 summarizes the final concentration for the primers, MgCl2, Dntps, Taq polymerase and the amplification program used for the molecular detection of Giardia duodenalis, Entamoeba histolytica/E. dispar, Cryptosporidium spp., Cyclospora spp. or Eimeria spp. and for Blastocystis spp.

Table 1.
Summary of the employed molecular techniques, the gene to be amplified, the species of parasites to be detected, the names of the primers and their sequence, the size of the amplified ones, the restriction enzymes and the bibliographic references of each test.

Table 2.
Summary of the final concentration of the primers, MgCl2, Dntps, Taq polymerase and the amplification program used for the molecular detection of Giardia duodenalis, Entamoeba histolytica/E. dispar, Cryptosporidium spp., Cyclospora spp., Eimeria spp., Blastocystis spp. and the bibliographic references for each test.

Results

To amplify the gene encoding Giardia duodenalis glutamate dehydrogenase (gdh) of approximately 432 bp, a semi-nested PCR was performed. Under the same conditions of amplification, a DNA segment of 343 ± 24 bp was amplified, which coincided with the 318 bp reported in the literature [2121. Read CM, Monis PT, Thompson RC. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol. 2004 Jun;4(2):125-30.], (Figure 1). It was possible to amplify DNA 1 fg/µL which was equivalent to 10 fg/rx (Figure 1A; Table 3) and 100 Q/mL (Figure 1B; Table 3). Figure 1C shows the amplification of the G. duodenalis gdh gene from different animal and human stool samples donated to Basic Research Laboratory. The DNA concentration range in these samples was found to be between 0.1 to 4 ng/µL of DNA and between 1,500 to 240,000 Q/mL. Figure 1D shows the RFLP obtained from the feces of a dog. RFLP is compatible with a BIII genotype, since fragment polymorphisms were obtained after digestion with Nla IV of 283 bp, 138 bp and 57 bp and with Rsa I of 310 bp, 137 bp and 36 bp [2020. Sepahvand A, Pestehchian N, Yousefi HA, Gharehbaba RP. Comparison and evaluation of four methods for extracting DNA from Giardia duodenalis cysts for PCR targeting the tpi gene. J Parasit Dis. 2017 Mar;41(1):263-7.

21. Read CM, Monis PT, Thompson RC. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol. 2004 Jun;4(2):125-30.

22. Pestehchian N, Rasekh H, Babaei Z, Yousefi HA, Eskandarian AA, Kazemi M, Akbari M. Identification of genotypes of Giardia duodenalis human isolates in Isfahan, Iran, using polymerase chain reaction - Restriction Fragment Length polymorphism. Adv Biomed Res. 2012 Dec 28;1:84.
-2323. Hooshyar H, Ghafarinasab S, Arbabi M, Delavari M, Rasti S. Genetic Variation of Giardia lamblia Isolates from Food-handlers in Kashan, Central Iran. Iran J Parasitol. 2017 Jan-Mar;12(1):83-9.].

Figure 1.
Detection and molecular characterization of the gene encoding G. duodenalis glutamate dehydrogenase (gdh) by means of nested semi-PCR and RFLP. (A) DNA detection limit (sensibility A). Lanes: (1) mix control; (2) 1 ng/µL DNA; (3) 0.1 ng/µL DNA; (4) 0.01 ng/µL DNA; (5) 1 pg/µL DNA; (6) 0.1 pg/µL DNA; (7) 0.01 pg/µL DNA; (8) 1 fg/µL DNA. (B) Cysts (Q) detection limit (sensibility B). Lanes: (1) 1 x 105 Q; (2) 1 x 104 Q; (3) 1 x 103 Q; (4) 1 x 102 Q; (5) 1 x 101 Q; (6) 1 x 100 Q; (7) 1 x 10-1 Q; (8) 1 x 10-2 Q; (9) mix control. (C) Molecular detection of the G. duodenalis gdh gene in human or animal samples. Lanes: (1) #33h (1 ng/µL; 2.4 x 105 Q/mL); (2) #33h (0.1 ng/µL); (3) #19d (3.7 ng/µL; 7500 Q/mL); (4) #22d (0.19 ng/µL; 12500 Q/mL); (5) #25d (1.98 ng/µL; 1481 Q/mL); (6) #26d (1.72 ng/µL, uncounted); (7) #24d (0.1 ng/µL; 2222 Q/mL); (8) mix control. (D) Molecular characterization by RFLP. Lanes: (1) digestion with Nla IV; (2) digestion with Rsa I. Bp: 50 bp molecular marker (#SM1103; GeneRuler™, ThermoScientific, USA); Bp*: 75 bp molecular marker (#SM1113; GeneRuler™, ThermoScientific, USA). h: human; d: dog. Arrows indicate the limit of detection.

The molecular detection of the genus Entamoeba spp. and the species of E. histolytica and E. dispar was carried out through a nested PCR, which amplified the gene that encoded the 16S-like rRNA, as described by Khairnar and Parija [2424. Khairnar K, Parija SC. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol. 2007 May 24;7:47.]. The external PCR that detected the genus of Entamoeba spp. performed with primers EG-1F and EG-2R, gave an amplification of approximately 887-898 bp (Figure 2A). Two nested PCRs were performed, one that detected E. histolytica with an amplification of 439 bp, with the primers EH-1F and EH-2R (Table 1), and another that detected E. dispar with an amplified 174 bp with the primers ED-1F and and ED-2R (Table 1). Nested PCR for E. dispar and E. histolytica resulted in amplification of 180 ± 6.5 bp (Figure 2B) and 386 ± 16.9 bp (Figure 2c), respectively, according to the literature [2424. Khairnar K, Parija SC. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol. 2007 May 24;7:47.]. According to these described conditions, it was possible to amplify 5 pg/µL of DNA or 12.5 pg DNA/rx PCR (sensitivity A), both for the nested PCR for E. dispar (Figure 2D) and for E. histolytica (Figure 2E) and up to 500 cysts/mL (sensitivity B) (Figure 2F; Table 3). It was possible to detect the presence of other species of the genus Entamoeba spp., amplified with the genus PCR, such as Entamoeba coli (Figure 2H).

Figure 2.
Molecular detection of the genus Entamoeba spp. and species E. dispar and E. histolytica by nested PCR. (A) PCR of the genus Entamoeba spp. (B) Nested PCR for E. dispar. (C) Nested PCR for E. histolytica. Lanes: (1) HL-67 (219 ng/µL); (2) HL-67 (21.9 ng/µL); (3) HL-68 (64.7 ng/µL); (4) HL-68 (6.47 ng/µL); (5) HL-27 (0.5 ng/µL); (6) HL-27 (0.05 ng/µL); (7) mix control. (D) Sensibility A for nested PCR to E. dispar. (E) Sensibility A for nested PCR to E. histolytica. Lanes: (1) 0.5 ng/µL DNA; (2) 0.05 ng/µL DNA; (3) 5 pg/µL DNA; (4) 0.5 pg/µL DNA; (5) 0.05 pg/µL DNA; (6) 5 fg/µL DNA; (7) 0.5 fg/µL DNA; (8) 0.05 fg/µL; (9) 1 atg/µL DNA; (10) mix control. (F) Cysts detection limit (sensibility B) for nested PCR E. dispar. (G) Cysts detection limit (sensibility B) for nested PCR E. dispar. Lanes: (1) 5000 Q; (2) 500 Q; (3) 50 Q; (4) 5 Q; (5) 0.5 Q; (6) 0.05 Q; (7) mix control; (8) positive control. (H) Nested PCR to E. dispar and E. histolytica. Lanes: (1) mixing control; (2) E. coli cysts (HL #35; 1.7 ng/µL); (3) E. coli cysts (HL #35; 0.17 ng/µL); (4-7) cysts of E. histolytica and E. dispar - lane (4) HL #27 (0.5 ng/µLDNA); (5) 0.05 ng/µL; (6) HL #27 (1.86 ng/µL); (7) HL #27 (0.186 ng/µL). Bp: 50 bp molecular marker (#SM1103; GeneRuler™, ThermoScientific, USA). Bp*: 100 Pb marker (#SM0323; Thermofisher, USA). HL: washed human feces. Arrows indicate the limit of detection.

To detect Cryptosporidium spp. and molecular characterization of Cryptosporidium parvum genotypes, a nested PCR was developed for the molecular detection of the genus by detecting the SSU 18S rRNA and RFLP gene for the identification of different genotypes of C. parvum, as published by Xiao et al. [2525. Xiao L, Morgan UM, Limor J, Escalante A, Arrowood M, Shulaw W, Thompson RC, Fayer R, Lal AA. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl Environ Microbiol. 1999 Aug;65(8):3386-91.]. The amplification conditions (Table 2) were identical to those recommended in the literature [2525. Xiao L, Morgan UM, Limor J, Escalante A, Arrowood M, Shulaw W, Thompson RC, Fayer R, Lal AA. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl Environ Microbiol. 1999 Aug;65(8):3386-91.], and allowed to amplify a DNA segment for the genus of 1359 ± 73 bp (Figure 3A) and 762 ± 38.3 bp for C. parvum (Figure 3B). Serial 10-fold dilutions of a DNA sample resulted in a nested PCR sensitivity for genus of 10 fg/µL that was equivalent to 50 fg DNA/rx (Figures 3A and 3B; Table 3). Figure 4C shows the molecular detection of Cryptosporidium spp. in a human stool sample demonstrating an increase in the level of detection in internal PCR at lower dilutions of DNA. Restriction enzyme digestion from nested PCR amplicon resulted in a polymorphism of 444 bp, 272 bp, and 130 bp after digestion with Ssp I, and 629 bp and 125 bp after digestion with Vsp I (Figure 3D), which coincided with C. parvum Bovine genotype B according to the literature [2525. Xiao L, Morgan UM, Limor J, Escalante A, Arrowood M, Shulaw W, Thompson RC, Fayer R, Lal AA. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl Environ Microbiol. 1999 Aug;65(8):3386-91.].

Figure 3.
Detection and molecular characterization of the genus of Cryptosporidium spp. (A) External PCR: amplification of a 1460 ± 135.2 bp DNA segment. (B) Internal PCR: amplification of a DNA segment of 762 ± 38.3 bp. Lanes: (1) 10 ng/µL; (2) 1 ng/µL; (3) 0.1 ng/µL; (4) 0.01 ng/µL; (5) 1 pg/µL; (6) 0.1 pg/µL; (7) 0.01 pg/µL; (8) 1 fg/µL; (9) 0.1 fg/µL; (10) mix control. (C) External PCR (R1) and internal PCR (R2), sample human feces. Lanes (1-3): pure DNA (1.64 ng/µL), diluted 1/10 and 1/100, respectively. Lanes 4-6: pure DNA (3.61 ng/µL), diluted 1/10 and 1/100, respectively. Lane (7) mix control. (D) Molecular characterization of genotype B bovine C. parvum by RFLP with restriction enzymes Ssp I and Vsp I. Lanes (1) digestion reaction; (2) mix control. Bp: 50 bp molecular marker (#SM1103; GeneRuler™, ThermoScientific, USA); Bp3: 100 bp molecular marker (#SM0323; ThermoScientific, USA). Arrows indicate the limit of detection.

The differential molecular detection between the genus of Cyclospora spp. and Eimeria spp. was carried out through a nested PCR by detecting the SSU 18S rRNA gene and RFLP with the restriction enzyme MnI I, according to what was published by Orlandi et al. [2626. Orlandi PA, Carter L, Brinker AM, da Silva AJ, Chu DM, Lampel KA, Monday SR. Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR. Appl Environ Microbiol. 2003 Aug;69(8):4806-13.]. The external amplification (R1) was performed with the pair of primers F1E and R2B and the internal one with F3E and R4B (Table 1). According to the amplification conditions for nested PCR (Table 2), a 284 ± 7.1 bp DNA fragment was amplified (Figure 4A), according to the literature [2626. Orlandi PA, Carter L, Brinker AM, da Silva AJ, Chu DM, Lampel KA, Monday SR. Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR. Appl Environ Microbiol. 2003 Aug;69(8):4806-13.], with a sensitivity A of 0.045 ng/µL or 225 pg/rx PCR (Figure 4A, Table 3), sensitivity B of 1000 oocysts (Figure 4B, Table 3). The nested PCR indiscriminately amplifies the genus of Cyclospora spp. and Eimeria spp. (Figure 4C) which may be different after performing the RFLP. The amplifications obtained were digested with MnI I, which allowed for the differentiation of the genus of Cyclospora spp. from the genus Eimeria spp. Three segments were obtained for the genus of Cyclospora spp. (133 bp, 104 bp, and 42 bp) and for Eimeria spp. (123 bp, 106 bp, and 61 bp) (Figure 4D), which coincided with the literature [2626. Orlandi PA, Carter L, Brinker AM, da Silva AJ, Chu DM, Lampel KA, Monday SR. Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR. Appl Environ Microbiol. 2003 Aug;69(8):4806-13.].

Figure 4.
Detection and molecular characterization of the genus Cyclospora spp. or Emeria spp. (A) Standardization of nested PCR using 2.5 µL (*) and 5 µL (**) of external PCR with a human HL sample with Cyclospora cayetanensis. Lanes: (1) mix control; (2) 4.5 ng/µL; (3) 0.45 ng/µL; (4) 0.045 ng/µL; (5) 4.5 pg/µL; (6) 0.45 pg/µL. (B) Qocysts (Q) detection limit (sensibility B). Lanes: (1) 7,5 x 103 Ooq.; (2) 1 x 103 Ooq.; (3) 1 x 102 Ooq.; (4) 1 x 101 Ooq.; (5) 1 x 100 Ooq.; (6) 1 x 10-1 Ooq.; (7) 1 x 10-2 Ooq.; (8) 1 x 10-3 Ooq.; (9) mix control; (10) positive control (human feces with Cyclospora oocysts). (C) Nested PCR (R2). Lanes: (1) mix control; (2) and (3) pure DNA (0.276 ng/µL) and diluted 1/10 of a sample of chicken feces with Eimeria spp., respectively; (4) and (5) pure DNA (1.64 ng/µL) and diluted 1/10 of a human stool sample with C. cayetanensis, respectively. (D) RFLP made with the enzyme MnI I. Lanes (1) and (2) Eimeria spp. RFLP; (3) and (4) Cyclospora spp. RFLP. Bp: 75 bp marker (#SM1331; Thermofisher, USA). Bp*: 50 bp molecular marker (#SM1103; GeneRuler™, ThermoScientific, USA). Arrows indicate the limit of detection.

The detection and molecular characterization for Blastocystis spp., was carried out following the amplification conditions by detecting the SSU 18S rRNA gene, described by Yoshikawa et al. [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.], were summarized in Table 1 and 2. With this protocol, a 1780 bp DNA fragment was amplified detecting the SSU 18S rRNA, which was consistent with the literature [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.]. This protocol made it possible to amplify DNA from washed stool pellets (HL) or washed pellets (CL) from stool cultures in MBDM. The PCR for Blastocystis spp. had a detection limit of 160 fg/µL of DNA (sensitivity A), equivalent to 800 fg of DNA/RX PCR (Figure 5A; Table 3), and a sensitivity B of 3600 vegetative forms (Figure 5C; Table 3). To increase the sensitivity of PCR, nested PCR was performed that amplified a 310 bp DNA segment that detects the SSU 18S rRNA, described by Stensvold et al. [2828. Stensvold R, Brillowska-Dabrowska A, Nielsen HV, Arendrup MC. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J Parasitol. 2006 Oct;92(5):1081-7.]. This nested PCR increased the sensitivity by 2 orders of magnitude to 1.6 fg/µL of DNA (sensitivity A), which was equivalent to 8 fg DNA/rx PCR, (Figure 5B; Table 3) and the detection of approximately 4 vegetative forms of the parasite, which corresponded to 3 orders of magnitude lower (sensitivity B) (Figure 5D, Table 3).

Figure 5.
Molecular detection of Blastocystis spp. (A) DNA detection limit for PCR 1780 bp. (B) DNA detection limit for nested PCR 310 bp. Lanes: (1) mix control; (2) 0.16 ng/µL; (3) 0.016 ng/µL; (4) 1.6 pg/µL; (5) 0.16 pg/µL; (6) 0.016 pg/µL; (7) 1.6 fg/µL; (8) 0.16 fg/µL; (9) 0.016 fg/µL. Bp: 100 bp molecular marker (#SM0323; Thermofisher, USA). (C) Detection of vegetative forms (vf) of Blastocystis for PCR of 1780 bp. (D) Detection of vf of Blastocystis for nested PCR 310 bp. Lanes: (1) 3.6 x 105 vf; (2) 3.6 x 104 vf; (3) 3.6 x 103 vf; (4) 3.6 x 102 vf; (5) 3.6 x 101 vf; (6) 3.6 x 100 vf; (7) 3.6 x 10-1 vf; (8) mix control; (9) positive control (CL-MMC). Bp: 50 bp molecular marker (#SM1103; GeneRuler™, ThermoScientific, USA). Arrows indicate the limit of detection.

The molecular characterization for Blastocystis spp. was carried out by RFLP the enzymes Rsa I and Hinf I, according to the suggestions of Yoshikawa et al. [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.]. These tests made it possible to discriminate the main genotypes that infect man (genotypes 1, 2, 3 and 4). The RFLPs obtained with each digestion were compared with those published by Yoshikawa et al. [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.]. Figure 6A shows the PCR amplification of 1780 bp from DNA extracted from human or pig stool samples (HL or CL) and in Figure 6B shows the RFLPs for subtypes (St) 1, 3 and 4 can be observed, whose polymorphism coincides with that published by Yoshikawa et al. [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.]. Of the 7 samples processed, some of them in duplicate (HL and CL), a polymorphism for St1 and St3 was obtained from human stool samples and a polymorphism for St4 for samples from pigs.

Figure 6.
Molecular characterization of Blastocystis spp. by PCR-RFLP. (A) PCR amplification of 1780 Bp from human (h) and pig (p) from HL (washed feces) or Boeck culture washed pellet (CL). (B) RFLP and subtypes (Sts) obtained with the enzymes Hinf I and Rsa I. Lanes: (1) control mix; (2) HL-11 (h; St3); (3) HL-12 (h); (4) CL-15 (p; St4); (5) CL-13 (h; St1); (6) CL-31 (h; St1); (7) CL-32 (h; St1); (8) CL-36 (p; St4); (9) CL-12 (h; St1); (10) CL-11 (h; St3). Bp*: 75 bp molecular marker (#SM1113¸ Thermofisher, USA); Bp: 50 bp molecular marker (#SM1103; GeneRuler™, ThermoScientific, USA).

The Table 3 summarizes the sensitivity of the standardized molecular techniques, in terms of the minimum concentration used to obtain a good performance amplified (sensitivity A), and in terms of the minimum number of vegetative forms, cysts or oocysts of protozoa detected by the molecular techniques used (sensitivity B).

Table 3.
Summary of the sensitivity of the different molecular techniques used to standardize the molecular detection of the parasites.

Discussion

Parasitological investigations of stool samples, in drinking water or food consumed raw or undercooked food, are widely used strategies in the research of parasitic infections in humans and animals or for investigation of infection routes [2929. Stark D, van Hal S, Marriott D, Ellis J, Harkness J. Irritable bowel syndrome: a review on the role of intestinal protozoa and the importance of their detection and diagnosis. Int J Parasitol. 2007 Jan;37(1):11-20.]. The investigation of the different stages of protozoa or chromist through molecular biology techniques based on the investigation of the detection and characterization of parasitic DNA [3030. Robertson LJ, Clark CG, Debenham JJ, Dubey JP, Kváč M, Li J, Ponce-Gordo F, Ryan U, Schares G, Su C, Tsaousis AD. Are molecular tools clarifying or confusing our understanding of the public health threat from zoonotic enteric protozoa in wildlife? Int J Parasitol Parasites Wildl. 2019 Feb 13;9:323-41.], from different matrices (water, feces, meat, soil), would improve the sensitivity and increase the probability of detection due to its high sensitivity and specificity [99. Jerez Puebla LE, Núñez-Fernández FA, Fraga Nodarse J, Atencio Millán I, Cruz Rodríguez I, Martínez Silva I, Ayllón Valdés L, Robertson LJ. Diagnosis of intestinal protozoan infections in patients in Cuba by microscopy and molecular methods: advantages and disadvantages. J Microbiol Methods. 2020 Dec;179:106102.]. In addition, it solves speciation problems, especially for those parasites species that are morphologically indistinguishable, as is the example for Cryptosporidium spp. [3131. Saki J, Foroutan-Rad M, Asadpouri R. Molecular Characterization of Cryptosporidium spp. in Wild Rodents of Southwestern Iran Using 18s rRNA Gene Nested-PCR-RFLP and Sequencing Techniques. J Trop Med. 2016;2016:6834206.] or Blastocystis spp. [3232. Khademvatan S, Masjedizadeh R, Yousefi-Razin E, Mahbodfar H, Rahim F, Yousefi E, Foroutan M. PCR-based molecular characterization of Blastocystis hominis subtypes in southwest of Iran. J Infect Public Health. 2018 Jan-Feb;11(1):43-7.], whose species or genotypes would be impossible to differentiate through the parasitic diagnosis. Besides, some subtypes, assemblages or genotypes of Blastocystis, Giardia duodenalis, Cryptosporidium spp., could be present in animals [3333. Alho AM, Lima C, Colella V, Madeira de Carvalho L, Otranto D, Cardoso L. Awareness of zoonotic diseases and parasite control practices: a survey of dog and cat owners in Qatar. Parasit Vectors. 2018 Mar 20;11(1):133.]. Therefore, the detection and molecular characterization of these protozoa, which can be found in animals or in environments and waters contaminated with animal feces, so that the molecular investigation of these protozoa could collaborate with the molecular epidemiology of protozoa with zoonotic potential [3434. Villamizar X, Higuera A, Herrera G, Vasquez-A LR, Buitron L, Muñoz LM, Gonzalez-C FE, Lopez MC, Giraldo JC, Ramírez JD. Molecular and descriptive epidemiology of intestinal protozoan parasites of children and their pets in Cauca, Colombia: a cross-sectional study. BMC Infect Dis. 2019 Feb 26;19(1):190.].

In this work, molecular techniques were standardized and tested to determine human pathogens, protozoa, and chromist. For all standardized molecular techniques (PCR, nested or semi-nested PCR, PCR-RFLP), the sensitivity of the technique was given in terms of the minimum quantity of DNA per µL or per PCR reaction (ADN/rx). The minimal quantity of biological forms detected by molecular tests, was called sensitivity B. Standardized tests allowed to find the species of the same genus, indistinguishable by microscopy, as is the case of the differentiation between E. histolytica and E. dispar, or arrive at genotypes (Blastocystis spp.) or assemblages (G. duodenalis), important for applying transition studies or molecular epidemiology.

The molecular detection of G. duodenalis in vegetable and fruit sediments has been reported through different methodologies, by direct microscopy via lugol [3535. Monge R, Arias ML. [Presence of various pathogenic microorganisms in fresh vegetables in Costa Rica]. Arch Latinoam Nutr. 1996 Dic; 46(4): 292-4. [Article in Spanish].], by fluorescence [3636. Utaaker KS, Kumar A, Joshi H, Chaudhary S, Robertson LJ. Checking the detail in retail: Occurrence of Cryptosporidium and Giardia on vegetables sold across different counters in Chandigarh, India. Int J Food Microbiol. 2017 Dec 18;263:1-8.] or by molecular biology techniques [3737. Colli CM, Bezagio RC, Nishi L, Bignotto TS, Ferreira ÉC, Falavigna-Guilherme AL, Gomes ML. Identical assemblage of Giardia duodenalis in humans, animals and vegetables in an urban area in southern Brazil indicates a relationship among them. PLoS One. 2015 Mar 11;10(3):e0118065.]. However, only molecular biology techniques can differientiate the genotype to which G. duodenalis belongs, which allows for determining the source (animal or human) of its origin. In this work, a semi-nested PCR for the molecular detection of G. duodenalis was used, reporting a sensitivity of one (1) fg/µL of DNA, which was 2000 times lower than that reported by Read et al. [2121. Read CM, Monis PT, Thompson RC. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol. 2004 Jun;4(2):125-30.]. When evaluating the detection limit of the forms of resistance, a detection limit of 100 cyst/rx was obtained for G. duodenalis, which was 100 times lower than that described by Read et al. [2121. Read CM, Monis PT, Thompson RC. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol. 2004 Jun;4(2):125-30.], and detected one trophozoite of G. duodenalis/rx. This is probably due to the use of pure cultures of G. duodenalis trophozoite cultures [2121. Read CM, Monis PT, Thompson RC. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol. 2004 Jun;4(2):125-30.]. Moreover, the use of stool samples and the presence of interferents can influence over PCR sensibility´s [3838. Perry GH, Marioni JC, Melsted P, Gilad Y. Genomic-scale capture and sequencing of endogenous DNA from feces. Mol Ecol. 2010 Dec;19(24):5332-44.], although the samples used in this study were previously washed. In addition, the extraction of DNA from the cyst has a greater degree of difficulty, due to the presence of constituent chitin of the wall of the cyst [3939. Temesgen TT, Barlaam A, Tysnes KR, Robertson LJ. Comparative evaluation of UNEX-based DNA extraction for molecular detection of Cyclospora cayetanensis, Toxoplasma gondii, and Cryptosporidium parvum as contaminants of berries. Food Microbiol. 2020 Aug;89:103447.]. Therefore, genotypes or assemblages of G. duodenalis research, using biomolecular technologies, would allow to make an inference of the parasite, which varies between hosts [4040. Monis PT, Andrews RH, Mayrhofer G, Ey PL. Genetic diversity within the morphological species Giardia intestinalis and its relationship to host origin. Infect Genet Evol. 2003 May;3(1):29-38., 4141. Feng Y, Xiao L. Zoonotic potential and molecular epidemiology of Giardia species and giardiasis. Clin Microbiol Rev. 2011 Jan;24(1):110-40.] and between geographic areas [3737. Colli CM, Bezagio RC, Nishi L, Bignotto TS, Ferreira ÉC, Falavigna-Guilherme AL, Gomes ML. Identical assemblage of Giardia duodenalis in humans, animals and vegetables in an urban area in southern Brazil indicates a relationship among them. PLoS One. 2015 Mar 11;10(3):e0118065.]. In this work, the detections of a BIII genotype was possible, using the RFLP from DNA amplified by a semi-nested PCR after digestion with endonuclease Nla IV and Rsa I, whose polymorphism coincided with other reports [2121. Read CM, Monis PT, Thompson RC. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol. 2004 Jun;4(2):125-30.

22. Pestehchian N, Rasekh H, Babaei Z, Yousefi HA, Eskandarian AA, Kazemi M, Akbari M. Identification of genotypes of Giardia duodenalis human isolates in Isfahan, Iran, using polymerase chain reaction - Restriction Fragment Length polymorphism. Adv Biomed Res. 2012 Dec 28;1:84.
-2323. Hooshyar H, Ghafarinasab S, Arbabi M, Delavari M, Rasti S. Genetic Variation of Giardia lamblia Isolates from Food-handlers in Kashan, Central Iran. Iran J Parasitol. 2017 Jan-Mar;12(1):83-9.].

The cysts of Entamoeba spp. can be detected with light microscopy depending on their morphological characteristics [4242. Shahnazi M, Jafari-Sabet M. Prevalence of parasitic contamination of raw vegetables in villages of Qazvin Province, Iran. Foodborne Pathog Dis. 2010 Sep;7(9):1025-30., 4343. Bekele F, Shumbej T. Fruit and vegetable contamination with medically important helminths and protozoans in Tarcha town, Dawuro zone, South West Ethiopia. Res Rep Trop Med. 2019 Apr 30;10:19-23.], with the help of Lugol [4444. Duedu KO, Yarnie EA, Tetteh-Quarcoo PB, Attah SK, Donkor ES, Ayeh-Kumi PF. A comparative survey of the prevalence of human parasites found in fresh vegetables sold in supermarkets and open-aired markets in Accra, Ghana. BMC Res Notes. 2014 Nov 25;7:836., 4545. Alemu G, Mama M, Misker D, Haftu D. Parasitic contamination of vegetables marketed in Arba Minch town, southern Ethiopia. BMC Infect Dis. 2019 May;19(1):410.]. Entamoeba histolytica (pathogenic amoeba) cysts are morphologically indistinguishable from E. dispar (commensal amoeba) and E. moshkovskii, considered free-living until it was isolated from a resident of Laredo, Texas, [4646. Clark CG, Diamond LS. The Laredo strain and other 'Entamoeba histolytica-like' amoebae are Entamoeba moshkovskii. Mol Biochem Parasitol. 1991 May;46(1):11-8.], who presented weight loss, epigastric pain and diarrhea. Other molecular epidemiological studies, through that the based on the determination of ribosomal SSU similar to 16S and the use of a multiplex PCR according to Khairnar and Parija [2424. Khairnar K, Parija SC. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol. 2007 May 24;7:47.], have revealed the presence of E. moshkovskii as responsible for gastrointestinal symptoms [4747. Hamzah Z, Petmitr S, Mungthin M, Leelayoova S, Chavalitshewinkoon-Petmitr P. Differential detection of Entamoeba histolytica, Entamoeba dispar, and Entamoeba moshkovskii by a single-round PCR assay. J Clin Microbiol. 2006 Sep;44(9):3196-200.

48. Tanyuksel M, Ulukanligil M, Guclu Z, Araz E, Koru O, Petri WA Jr. Two cases of rarely recognized infection with Entamoeba moshkovskii. Am J Trop Med Hyg. 2007 Apr;76(4):723-4.
-4949. Ayed SB, Aoun K, Maamouri N, Abdallah RB, Bouratbine A. First molecular identification of Entamoeba moshkovskii in human stool samples in Tunisia. Am J Trop Med Hyg. 2008 Nov;79(5):706-7.]. Therefore, determining the species is a critical step for the establishment of a treatment, when the pathogen is detected in human feces samples, or for indicating the origin of the contamination, if it is detected in vegetables or fruits for human consumption. In this study, nested PCR for the detection of E. histolytica and E. dispar was able to detect 500 cysts in washed stool samples (LH), well above that reported by Khairnar and Parija [2424. Khairnar K, Parija SC. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol. 2007 May 24;7:47.]. In comparison with our study, Khairnar and Parija [2424. Khairnar K, Parija SC. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol. 2007 May 24;7:47.] performed the experiments with trophozoite cultures, enriched and free from the remains of the stool samples. The nested multiplex PCR detection limit for E. histolytica, E. dispar and E. moshkovskii was found to be approximately 25 cells of Entamoeba protozoa, since 2.5 μL of template DNA (1000 parasites/100 μL of TE buffer), so it was expected that in our work the sensitivity B of the tests would be lower. On the other hand, the use of stool samples with cysts as a DNA extraction matrix can inhibit PCR by having a high amount of bacteria and detritus typical of digestion [3838. Perry GH, Marioni JC, Melsted P, Gilad Y. Genomic-scale capture and sequencing of endogenous DNA from feces. Mol Ecol. 2010 Dec;19(24):5332-44.] and the cyst, with respect to the trophozoite, has a DNA extraction difficulty inherent in its conformation, such as a resistance structure [3939. Temesgen TT, Barlaam A, Tysnes KR, Robertson LJ. Comparative evaluation of UNEX-based DNA extraction for molecular detection of Cyclospora cayetanensis, Toxoplasma gondii, and Cryptosporidium parvum as contaminants of berries. Food Microbiol. 2020 Aug;89:103447.].

Cryptosporidium spp. are protozoan parasites that infect humans and animals, and the second most common cause of diarrhea in children after rotavirus [5050. Bouzid M, Kintz E, Hunter PR. Risk factors for Cryptosporidium infection in low and middle income countries: A systematic review and meta-analysis. PLoS Negl Trop Dis. 2018 Jun 7;12(6):e0006553.]. Cryptosporidium spp. it is characterized by its extensive genetic variation that results in the existence of 38 species and more than 60 genotypes of this parasite [5151. Feng Y, Ryan UM, Xiao L. Genetic Diversity and Population Structure of Cryptosporidium. Trends Parasitol. 2018 Nov;34(11):997-1011.]. At least 20 different species cause moderate or severe infections in humans, of which C. hominis and C. parvum are the main causative agents [5252. Khan A, Shaik JS, Grigg ME. Genomics and molecular epidemiology of Cryptosporidium species. Acta Trop. 2018 Aug;184:1-14.]. Molecular tools have been developed to detect and differentiate Cryptosporidium spp. at the species/genotype and subtype level. These tools have been used increasingly to characterize the transmission of Cryptosporidium spp. in humans and animals [5353. Xiao L. Molecular epidemiology of cryptosporidiosis: an update. Exp Parasitol. 2010 Jan;124(1):80-9.]. In addition, they have also been used to investigate the sources of infection for humans, such as in water collections [5454. Zahedi A, Monis P, Gofton AW, Oskam CL, Ball A, Bath A, Bartkow M, Robertson I, Ryan U. Cryptosporidium species and subtypes in animals inhabiting drinking water catchments in three states across Australia. Water Res. 2018 May 1;134:327-40., 5555. Hamdy D, El-Badry A, Abd El Wahab W. Assessment of Giardia and Cryptosporidium Assemblages/Species and Their Viability in Potable Tap Water in Beni-Suef, Egypt Using Nested PCR/RFLP and Staining. Iran J Parasitol. 2019 Jul-Sep;14(3):368-78.] and in vegetables and fruits for human consumption. Genotyping tools based on DNA sequences of antigens and housekeeping genes identified genotype 1 for the human genotype and genotype 2 for the bovine genotype, within the C. parvum umbrela, gave rise to C. hominis and C. parvum, respectively, both infectious for immunocompetent and immunosuppressed people [5656. Xiao L, Fayer R, Ryan U, Upton SJ. Cryptosporidium taxonomy: recent advances and implications for public health. Clin Microbiol Rev. 2004 Jan;17(1):72-97., 5757. Cacciò SM. Molecular epidemiology of human cryptosporidiosis. Parassitologia. 2005 Jun;47(2):185-92.].

In this work, it was possible to determine the sensitivity of nested PCR for the detection of 18S SSU rRNA for Cryptosporidium spp., at 10 fg/µL (equivalent to 50 fg DNA/rx). There is no literature reporting sensitivity A for nested PCR to detect this protozoan, but there are reports where this PCR allows amplifying 1 µL of DNA, without specifying the concentration of the DNA used [5858. Jiang J, Alderisio KA, Singh A, Xiao L. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl Environ Microbiol. 2005 Mar;71(3):1135-41. ]. In this study Cryptosporidium spp. oocysts were not quantified in this study, but it can be done in a Neubauer chamber after concentration by flotation in sucrose [5959. Toledo J, Lombardelli J, Galarza R, Tiranti K, Garro CJ, Florin-Christensen M, Schnittger L, Tomazic ML. Comparison of DNA extraction methods to improve the molecular diagnosis of Cryptosporidium spp. from fecal samples of calves. Rev Inv Cient Univ Morón. 2018 Jul;2(4):17-28. ] or molecularly, when performing a quantitative or real-time PCR (q-PCR), where the number of copies per oocyst can be estimated according to the gene under investigation. In this regard, Li et al. [6060. Li N, Neumann NF, Ruecker N, Alderisio KA, Sturbaum GD, Villegas EN, 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 Sep 1;81(17):5845-54.] standardized the detection of oocysts of different Cryptosporiridium species and concluded that the amount of these parasites is determined by the fact that the gene used as a target has 20 copies per oocyst. Therefore, if real-time PCR can detect at least 20 copies of the gene, the sensitivity of the molecular test would be one oocyst. Real-time PCR would be a more sensitive technique and could quantify the copy number for a parasite like Cryptosporidium spp., which would be more convenient [6161. Hadfield SJ, Robinson G, Elwin K, Chalmers RM. Detection and differentiation of Cryptosporidium spp. in human clinical samples by use of real-time PCR. J Clin Microbiol. 2011 Mar;49(3):918-24.

62. Mary C, Chapey E, Dutoit E, Guyot K, Hasseine L, Jeddi F, Menotti J, Paraud C, Pomares C, Rabodonirina M, Rieux A, Derouin F; ANOFEL Cryptosporidium National Network. 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 Aug;51(8):2556-63.
-6363. Rousseau A, La Carbona S, Dumètre A, Robertson LJ, Gargala G, Escotte-Binet S, Favennec L, Villena I, Gérard C, Aubert D. Assessing viability and infectivity of foodborne and waterborne stages (cysts/oocysts) of Giardia duodenalis, Cryptosporidium spp., and Toxoplasma gondii: a review of methods. Parasite. 2018;25:14.] than its detection after a flotation concentration in sucrose [5959. Toledo J, Lombardelli J, Galarza R, Tiranti K, Garro CJ, Florin-Christensen M, Schnittger L, Tomazic ML. Comparison of DNA extraction methods to improve the molecular diagnosis of Cryptosporidium spp. from fecal samples of calves. Rev Inv Cient Univ Morón. 2018 Jul;2(4):17-28. ].

Regarding the simultaneous molecular detection for the Cyclospora spp. and Eimeria spp., it is possible to discriminate Cyclospora cayetanensis by RFLP only if the sample tested is from human feces. The detection of the genus of Cyclospora spp., in another type of sample, implies the presence of other species of animal origin (non-human primates) that could be present in the analyzed samples of sediment of vegetables, fruits or drinking water [6464. Hussein EM. Molecular identification of Cyclospora spp. using multiplex PCR from diarrheic children compared to others conventional methods. J Egypt Soc Parasitol. 2007 Aug;37(2):585-98., 6565. Li W, Kiulia NM, Mwenda JM, Nyachieo A, Taylor MB, Zhang X, Xiao L. Cyclospora papionis, Cryptosporidium hominis, and human-pathogenic Enterocytozoon bieneusi in captive baboons in Kenya. J Clin Microbiol. 2011 Dec;49(12):4326-9.], and although these are not pathogenic for humans, it would illustrate the contamination of cultures with non-human feces. Therefore, the molecular detection recommended for Cyclospora spp. would be sequencing of conserved genes such as those of SSU rRNA, after amplification by PCR [6060. Li N, Neumann NF, Ruecker N, Alderisio KA, Sturbaum GD, Villegas EN, 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 Sep 1;81(17):5845-54.], or using more specific primers that prevent the amplification of DNA from oocysts of another genus [6666. Qvarnstrom Y, Benedict T, Marcet PL, Wiegand RE, Herwaldt BL, da Silva AJ. Molecular detection of Cyclospora cayetanensis in human stool specimens using UNEX-based DNA extraction and real-time PCR. Parasitology. 2018 Jun;145(7):865-870.]. On the other hand, finding Eimeria spp. in sediment samples of fruits or vegetables for human consumption could mean the use of chicken manure [6767. Mathis GF, McDougald LR. Drug responsiveness of field isolates of chicken Coccidia. Poult Sci. 1982 Jan;61(1):38-45.], bovine [6868. Cruvinel LB, Nicaretta JE, Bastos TSA, Couto LFM, Santos JBD, Zapa DMB, Cavalcante ASA, Cruz BC, Borges DGL, Borges FA, Soares VE, Lopes WDZ. Eimeria species in dairy and beef cattle of different ages in Goiás state, Brazil. Rev Bras Parasitol Vet. 2018 Apr-Jun;27(2):169-76.] or equine manure [6969. Dubey JP, Bauer C. A review of Eimeria infections in horses and other equids. Vet Parasitol. 2018 May 30;256:58-70.], between others, as compost to fertilize vegetable or fruit crops, or the use of water contaminated with animal feces. Therefore, it would be important to identify the species to be able to identify the origin of this protozoan.

In this study, the limit of sensitivity for the detection of DNA for the genus of Cyclospora spp. or Eimeria spp. was 225 pg/rx. The minimal amount of oocysts of Cyclospora spp. or Eimeria spp. 1000 oocysts/rx were found to be 10 times less than the 100 oocysts of Cyclospora spp. or Eimeria spp., reported by PCR by Orlandi et al. [2626. Orlandi PA, Carter L, Brinker AM, da Silva AJ, Chu DM, Lampel KA, Monday SR. Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR. Appl Environ Microbiol. 2003 Aug;69(8):4806-13.]. In this regard, the samples used by Orlandi et al. [2626. Orlandi PA, Carter L, Brinker AM, da Silva AJ, Chu DM, Lampel KA, Monday SR. Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR. Appl Environ Microbiol. 2003 Aug;69(8):4806-13.] were kept in 2% potassium dichromate, which favored oocyst maturation. This could favor the sensitivity of the applied molecular technique, since a mature oocyst will have a greater amount of DNA than inmature ones, facilitating its finding through molecular methodologies. In this context, and during the standardization of molecular techniques for the detection of Cryptosporidium spp. in calves, Toledo et al. [5959. Toledo J, Lombardelli J, Galarza R, Tiranti K, Garro CJ, Florin-Christensen M, Schnittger L, Tomazic ML. Comparison of DNA extraction methods to improve the molecular diagnosis of Cryptosporidium spp. from fecal samples of calves. Rev Inv Cient Univ Morón. 2018 Jul;2(4):17-28. ] demonstrated, that the amount of DNA required to detect the 18S rRNA gene [2525. Xiao L, Morgan UM, Limor J, Escalante A, Arrowood M, Shulaw W, Thompson RC, Fayer R, Lal AA. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl Environ Microbiol. 1999 Aug;65(8):3386-91.], can be 16 times lower, when using an enriched sample of oocysts stored in potassium dichromate (0.025 ng of DNA) than from faeces (0.4 ng of DNA). Therefore, the sensitivity of a PCR will depend on how enriched and pure it is, which supports the decrease in sensitivity in some of the tests carried out in this work compared to the literature.

Regarding the molecular detection limit for Blastocystis spp., a detection limit of 3600 evolutionary forms per reaction was obtained by amplifying the SSUrRNA 18S gene of 1780 bp by endpoint PCR, as described by Yoshikawa et al. [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.], but almost 4 evolutionary forms per reaction, when performing the nested PCR, whose endpoint PCR was described by Stensvold et al. [2828. Stensvold R, Brillowska-Dabrowska A, Nielsen HV, Arendrup MC. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J Parasitol. 2006 Oct;92(5):1081-7.]. When compared with references in the literature, Yoshikawa et al. [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.] do not mention the parasite molecular detection limit. Stensvold et al. [2828. Stensvold R, Brillowska-Dabrowska A, Nielsen HV, Arendrup MC. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J Parasitol. 2006 Oct;92(5):1081-7.] reported a PCR sensitivity of 80 parasites per gram of feces, which was 20 times higher than that reported in this work, after using the PCR described by them as a nested PCR, from the amplified yield obtained by amplifying the 1780 bp SSUrRNA gene described by Yoshikawa et al. [2727. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.]. By molecularly characterizing Blastocystis spp. a predominance of St3 over St1 and St4 was obtained, all of them found as infectious subtypes (Sts) in man [7070. Stensvold CR, Lewis HC, Hammerum AM, Porsbo LJ, Nielsen SS, Olsen KE, Arendrup MC, Nielsen HV, Mølbak K. Blastocystis: unravelling potential risk factors and clinical significance of a common but neglected parasite. Epidemiol Infect. 2009 Nov;137(11):1655-63.

71. Noradilah SA, Moktar N, Anuar TS, Lee IL, Salleh FM, Manap SNAA, Mohtar NSHM, Azrul SM, Abdullah WO, Nordin A, Abdullah SR. Molecular epidemiology of blastocystosis in Malaysia: does seasonal variation play an important role in determining the distribution and risk factors of Blastocystis subtype infections in the Aboriginal community? Parasit Vectors. 2017 Jul 31;10(1):360.
-7272. Alfellani MA, Stensvold CR, Vidal-Lapiedra A, Onuoha ES, Fagbenro-Beyioku AF, Clark CG. Variable geographic distribution of Blastocystis subtypes and its potential implications. Acta Trop. 2013 Apr;126(1):11-8.], and among the nine Sts present in man (St1 al St9) with St2 being the most common Sts found in human feces [7272. Alfellani MA, Stensvold CR, Vidal-Lapiedra A, Onuoha ES, Fagbenro-Beyioku AF, Clark CG. Variable geographic distribution of Blastocystis subtypes and its potential implications. Acta Trop. 2013 Apr;126(1):11-8.]. The RFLP performed allows defining the subtypes found in 95% of humans infected with Blastocystis [7171. Noradilah SA, Moktar N, Anuar TS, Lee IL, Salleh FM, Manap SNAA, Mohtar NSHM, Azrul SM, Abdullah WO, Nordin A, Abdullah SR. Molecular epidemiology of blastocystosis in Malaysia: does seasonal variation play an important role in determining the distribution and risk factors of Blastocystis subtype infections in the Aboriginal community? Parasit Vectors. 2017 Jul 31;10(1):360.]. Some of the Sts found have been related to gastrointestinal symptoms [7171. Noradilah SA, Moktar N, Anuar TS, Lee IL, Salleh FM, Manap SNAA, Mohtar NSHM, Azrul SM, Abdullah WO, Nordin A, Abdullah SR. Molecular epidemiology of blastocystosis in Malaysia: does seasonal variation play an important role in determining the distribution and risk factors of Blastocystis subtype infections in the Aboriginal community? Parasit Vectors. 2017 Jul 31;10(1):360.

72. Alfellani MA, Stensvold CR, Vidal-Lapiedra A, Onuoha ES, Fagbenro-Beyioku AF, Clark CG. Variable geographic distribution of Blastocystis subtypes and its potential implications. Acta Trop. 2013 Apr;126(1):11-8.
-7373. Tan KS, Mirza H, Teo JD, Wu B, Macary PA. Current Views on the Clinical Relevance of Blastocystis spp. Curr Infect Dis Rep. 2010 Jan;12(1):28-35.] and others to their absence [7474. El Safadi D, Cian A, Nourrisson C, Pereira B, Morelle C, Bastien P, Bellanger AP, Botterel F, Candolfi E, Desoubeaux G, Lachaud L, Morio F, Pomares C, Rabodonirina M, Wawrzyniak I, Delbac F, Gantois N, Certad G, Delhaes L, Poirier P, Viscogliosi E. Prevalence, risk factors for infection and subtype distribution of the intestinal parasite Blastocystis sp. from a large-scale multi-center study in France. BMC Infect Dis. 2016 Aug 26;16(1):451.] or as part of the intestinal microbiota [7575. Tito RY, Chaffron S, Caenepeel C, Lima-Mendez G, Wang J, Vieira-Silva S, Falony G, Hildebrand F, Darzi Y, Rymenans L, Verspecht C, Bork P, Vermeire S, Joossens M, Raes J. Population-level analysis of Blastocystis subtype prevalence and variation in the human gut microbiota. Gut. 2019 Jul;68(7):1180-9.].

Conclusions

The sensitivity A (minimum quantity of DNA) of the standardized molecular techniques by PCR reaction was 10 fg, 12.5 pg, 50 fg, 225 pg 800 fg and 8 fg for G. duodenalis (semi-nested PCR), Entamoeba spp. (genus), Cryptosporidium spp., for the simultaneous genus of Cyclospora spp. and Eimeria spp. and for Blastocystis spp. after performing 1780 bp PCR or, nested PCR (310 bp), respectively. For the parasites in which it was possible, the minimum number of protozoa or chromists that were detected by the molecular technique used was determined, which was 100 and 500 cysts for G. duodenalis and Entamoeba genus, of 1000 oocysts for the detection of forms of Cyclospora spp. or Eimeria spp. and 3600 or four vegetative ones for Blastocystis spp. (PCR 1780 bp) or Blastocystis spp. (nested PCR; 310 bp), respectively. The molecular detection of protozoa and chromist was achieved and the molecular characterization allowed the genotyping of some of the parasites such as Giardia duodenalis, Cryptosporidium spp., and Blastocystis spp. For the molecular detection of Cyclospora species, it would be advisable to sequence conserved genes such as those of the SSU rRNA, after their PCR amplification [5252. Khan A, Shaik JS, Grigg ME. Genomics and molecular epidemiology of Cryptosporidium species. Acta Trop. 2018 Aug;184:1-14.] or to use more specific primers for avoid amplification of oocysts’ DNA of another genus [5858. Jiang J, Alderisio KA, Singh A, Xiao L. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl Environ Microbiol. 2005 Mar;71(3):1135-41. ]. For the detection of Cryptosporidium spp. gene sequencing is recommended. This study opens the door to the molecular epidemiology of intestinal protozoosis and Blastocystis spp., which can be used for epidemiological studies in humans, animals, sources of transmission and as diagnostic tools in countries where intestinal parasites are a public health problem.

Acknowledgments

The authors are grateful to Mrs. Adriana Vethencourt for her collaboration in writing in native English and Mrs. Carmen Ramirez for her secretarial work.

References

  • 1. Chelkeba L, Mekonnen Z, Alemu Y, Emana D. Epidemiology of intestinal parasitic infections in preschool and school-aged Ethiopian children: a systematic review and meta-analysis. BMC Public Health. 2020 Jan 28;20(1):117.
  • 2. Costa Rica. Ministerio de Salud. Instituto Costarricense de Investigación y Enseñanza en Nutrición y Salud. Caja Costarricense de Seguro Social. Instituto Nacional de Estadística y Censos. Instituto Costarricense Sobre Drogas. Encuesta Nacional de Nutrición 2008-2009. Fascículo 5: Parásitos Intestinales. San José, Costa Rica: El Ministerio, 2012. Available from: https://www.binasss.sa.cr/opac-ms/media/digitales/Encuesta%20Nacional%20de%20Nutrici%C3%B3n.%20Fasc%C3%ADculo%205.%20Par%C3%A1sitos%20intestinales.pdf
    » https://www.binasss.sa.cr/opac-ms/media/digitales/Encuesta%20Nacional%20de%20Nutrici%C3%B3n.%20Fasc%C3%ADculo%205.%20Par%C3%A1sitos%20intestinales.pdf
  • 3. Mata L. Erradicación de helmintos intestinales en Costa Rica. Rev Med Hosp Nac Niños Costa Rica. 1985;20(2):151-64.
  • 4. Abrahams-Sandí E, Solano M., Rodríguez B. Prevalencia de parásitos intestinales en escolares de Limón Centro, Costa Rica. Rev Costarric Cienc Med. 2005;26(1-2):33-8.
  • 5. Arévalo M, Cortés X, Barrantes K, Achí R. Prevalence of intestinal parasitosis in children from the community of Los Cuadros, Goicoechea, Costa Rica. 2002-2003. Rev Costarric Cienc Med. 2007 Jun;28(1-2):37-45.
  • 6. Solano-Barquero M, Montero-Salguero A, León-Alán D, Santamaría-Ulloa C, Mora A M, Reyes-Lizano L. Prevalence of parasitosis in children aged 1 to 7 years in a vulnerable condition in the Central-South Region of Costa Rica. Acta Med Costarric. 2018 Apr-Jun;60(2):19-29.
  • 7. Solano L, Acuña I, Barón M, Morón DE SA, Sánchez, A. Intestinal parasitic infestation and others infectious backgrounds influence in the antropometric nutritional status of children in poverty. Parasitol Latinoam. 2008;63(1-4):12-9.
  • 8. Akgül Ö, Kart Yaşar K, Sapmaz B, Kırkoyun Uysal H, Yıldırmak T, Şimşek F, Karasakal ÖF, Çalışkan R, Öner YA. [Detection of intestinal parasites with conventional and molecular methods in follow-up HIV/AIDS cases]. Mikrobiyol Bul. 2018 Jul;52(3):273-83. [Article in Turkish].
  • 9. Jerez Puebla LE, Núñez-Fernández FA, Fraga Nodarse J, Atencio Millán I, Cruz Rodríguez I, Martínez Silva I, Ayllón Valdés L, Robertson LJ. Diagnosis of intestinal protozoan infections in patients in Cuba by microscopy and molecular methods: advantages and disadvantages. J Microbiol Methods. 2020 Dec;179:106102.
  • 10. Rivero de Rodríguez Z. Detection of Entamoeba moshkovskii in humans: a new diagnostic problem in amebiasis diagnosis. Kasmera. 2013;41(1):42-9.
  • 11. Salehi R, Haghighi A, Stensvold CR, Kheirandish F, Azargashb E, Raeghi S, Kohansal C, Bahrami F. Prevalence and subtype identification of Blastocystis isolated from humans in Ahvaz, Southwestern Iran. Gastroenterol Hepatol Bed Bench. 2017;10(3):235-41.
  • 12. Tahvildar-Biderouni F, Salehi N. Detection of Cryptosporidium infection by modified ziehl-neelsen and PCR methods in children with diarrheal samples in pediatric hospitals in Tehran. Gastroenterol Hepatol Bed Bench. 2014;7(2):125-30.
  • 13. Zahedi A, Ryan U. Cryptosporidium - An update with an emphasis on foodborne and waterborne transmission. Res Vet Sci. 2020 Oct;132:500-12.
  • 14. Feng Y, Ryan UM, Xiao L. Genetic Diversity and Population Structure of Cryptosporidium Trends Parasitol. 2018 Nov;34(11):997-1011.
  • 15. Xiao L, Feng Y. Molecular epidemiologic tools for waterborne pathogens Cryptosporidium spp. and Giardia duodenalis Food Waterborne Parasitol. 2017 Sep 29;8-9:14-32.
  • 16. Nader JL, Mathers TC, Ward BJ, Pachebat JA, Swain MT, Robinson G, Chalmers RM, Hunter PR, van Oosterhout C, Tyler KM. Evolutionary genomics of anthroponosis in Cryptosporidium Nat Microbiol. 2019 May;4(5):826-36.
  • 17. Guzmán de Rondón C, Vethencourt MA, Galindo M, Chacón N, Wagner C, Nessi A. Comportamiento biológico de Blastocystis hominis en pacientes tratados con Secnidazol (Unidazol®) / Biological behavior of Blastocystis hominis in patients treated with Secnidazol (Unidazol®). Rev Soc Venez Microbiol. 2008 Jun;28(1):66-71.
  • 18. Kageruka P, Brandt JR, Taelman H, Jonas C. Modified Köster staining method for the diagnosis of cryptosporidiosis. Ann Soc Belg Med Trop. 1984 Jun;64(2):171-5.
  • 19. Murray PR, Baron EJ. Manual of Clinical Microbiology. 9th ed. Murray PR, editor. Washington, D.C: ASM Press; 2007.
  • 20. Sepahvand A, Pestehchian N, Yousefi HA, Gharehbaba RP. Comparison and evaluation of four methods for extracting DNA from Giardia duodenalis cysts for PCR targeting the tpi gene. J Parasit Dis. 2017 Mar;41(1):263-7.
  • 21. Read CM, Monis PT, Thompson RC. Discrimination of all genotypes of Giardia duodenalis at the glutamate dehydrogenase locus using PCR-RFLP. Infect Genet Evol. 2004 Jun;4(2):125-30.
  • 22. Pestehchian N, Rasekh H, Babaei Z, Yousefi HA, Eskandarian AA, Kazemi M, Akbari M. Identification of genotypes of Giardia duodenalis human isolates in Isfahan, Iran, using polymerase chain reaction - Restriction Fragment Length polymorphism. Adv Biomed Res. 2012 Dec 28;1:84.
  • 23. Hooshyar H, Ghafarinasab S, Arbabi M, Delavari M, Rasti S. Genetic Variation of Giardia lamblia Isolates from Food-handlers in Kashan, Central Iran. Iran J Parasitol. 2017 Jan-Mar;12(1):83-9.
  • 24. Khairnar K, Parija SC. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar DNA in stool samples. BMC Microbiol. 2007 May 24;7:47.
  • 25. Xiao L, Morgan UM, Limor J, Escalante A, Arrowood M, Shulaw W, Thompson RC, Fayer R, Lal AA. Genetic diversity within Cryptosporidium parvum and related Cryptosporidium species. Appl Environ Microbiol. 1999 Aug;65(8):3386-91.
  • 26. Orlandi PA, Carter L, Brinker AM, da Silva AJ, Chu DM, Lampel KA, Monday SR. Targeting single-nucleotide polymorphisms in the 18S rRNA gene to differentiate Cyclospora species from Eimeria species by multiplex PCR. Appl Environ Microbiol. 2003 Aug;69(8):4806-13.
  • 27. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol. 2000 Apr;38(4):1324-30.
  • 28. Stensvold R, Brillowska-Dabrowska A, Nielsen HV, Arendrup MC. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J Parasitol. 2006 Oct;92(5):1081-7.
  • 29. Stark D, van Hal S, Marriott D, Ellis J, Harkness J. Irritable bowel syndrome: a review on the role of intestinal protozoa and the importance of their detection and diagnosis. Int J Parasitol. 2007 Jan;37(1):11-20.
  • 30. Robertson LJ, Clark CG, Debenham JJ, Dubey JP, Kváč M, Li J, Ponce-Gordo F, Ryan U, Schares G, Su C, Tsaousis AD. Are molecular tools clarifying or confusing our understanding of the public health threat from zoonotic enteric protozoa in wildlife? Int J Parasitol Parasites Wildl. 2019 Feb 13;9:323-41.
  • 31. Saki J, Foroutan-Rad M, Asadpouri R. Molecular Characterization of Cryptosporidium spp. in Wild Rodents of Southwestern Iran Using 18s rRNA Gene Nested-PCR-RFLP and Sequencing Techniques. J Trop Med. 2016;2016:6834206.
  • 32. Khademvatan S, Masjedizadeh R, Yousefi-Razin E, Mahbodfar H, Rahim F, Yousefi E, Foroutan M. PCR-based molecular characterization of Blastocystis hominis subtypes in southwest of Iran. J Infect Public Health. 2018 Jan-Feb;11(1):43-7.
  • 33. Alho AM, Lima C, Colella V, Madeira de Carvalho L, Otranto D, Cardoso L. Awareness of zoonotic diseases and parasite control practices: a survey of dog and cat owners in Qatar. Parasit Vectors. 2018 Mar 20;11(1):133.
  • 34. Villamizar X, Higuera A, Herrera G, Vasquez-A LR, Buitron L, Muñoz LM, Gonzalez-C FE, Lopez MC, Giraldo JC, Ramírez JD. Molecular and descriptive epidemiology of intestinal protozoan parasites of children and their pets in Cauca, Colombia: a cross-sectional study. BMC Infect Dis. 2019 Feb 26;19(1):190.
  • 35. Monge R, Arias ML. [Presence of various pathogenic microorganisms in fresh vegetables in Costa Rica]. Arch Latinoam Nutr. 1996 Dic; 46(4): 292-4. [Article in Spanish].
  • 36. Utaaker KS, Kumar A, Joshi H, Chaudhary S, Robertson LJ. Checking the detail in retail: Occurrence of Cryptosporidium and Giardia on vegetables sold across different counters in Chandigarh, India. Int J Food Microbiol. 2017 Dec 18;263:1-8.
  • 37. Colli CM, Bezagio RC, Nishi L, Bignotto TS, Ferreira ÉC, Falavigna-Guilherme AL, Gomes ML. Identical assemblage of Giardia duodenalis in humans, animals and vegetables in an urban area in southern Brazil indicates a relationship among them. PLoS One. 2015 Mar 11;10(3):e0118065.
  • 38. Perry GH, Marioni JC, Melsted P, Gilad Y. Genomic-scale capture and sequencing of endogenous DNA from feces. Mol Ecol. 2010 Dec;19(24):5332-44.
  • 39. Temesgen TT, Barlaam A, Tysnes KR, Robertson LJ. Comparative evaluation of UNEX-based DNA extraction for molecular detection of Cyclospora cayetanensis, Toxoplasma gondii, and Cryptosporidium parvum as contaminants of berries. Food Microbiol. 2020 Aug;89:103447.
  • 40. Monis PT, Andrews RH, Mayrhofer G, Ey PL. Genetic diversity within the morphological species Giardia intestinalis and its relationship to host origin. Infect Genet Evol. 2003 May;3(1):29-38.
  • 41. Feng Y, Xiao L. Zoonotic potential and molecular epidemiology of Giardia species and giardiasis. Clin Microbiol Rev. 2011 Jan;24(1):110-40.
  • 42. Shahnazi M, Jafari-Sabet M. Prevalence of parasitic contamination of raw vegetables in villages of Qazvin Province, Iran. Foodborne Pathog Dis. 2010 Sep;7(9):1025-30.
  • 43. Bekele F, Shumbej T. Fruit and vegetable contamination with medically important helminths and protozoans in Tarcha town, Dawuro zone, South West Ethiopia. Res Rep Trop Med. 2019 Apr 30;10:19-23.
  • 44. Duedu KO, Yarnie EA, Tetteh-Quarcoo PB, Attah SK, Donkor ES, Ayeh-Kumi PF. A comparative survey of the prevalence of human parasites found in fresh vegetables sold in supermarkets and open-aired markets in Accra, Ghana. BMC Res Notes. 2014 Nov 25;7:836.
  • 45. Alemu G, Mama M, Misker D, Haftu D. Parasitic contamination of vegetables marketed in Arba Minch town, southern Ethiopia. BMC Infect Dis. 2019 May;19(1):410.
  • 46. Clark CG, Diamond LS. The Laredo strain and other 'Entamoeba histolytica-like' amoebae are Entamoeba moshkovskii Mol Biochem Parasitol. 1991 May;46(1):11-8.
  • 47. Hamzah Z, Petmitr S, Mungthin M, Leelayoova S, Chavalitshewinkoon-Petmitr P. Differential detection of Entamoeba histolytica, Entamoeba dispar, and Entamoeba moshkovskii by a single-round PCR assay. J Clin Microbiol. 2006 Sep;44(9):3196-200.
  • 48. Tanyuksel M, Ulukanligil M, Guclu Z, Araz E, Koru O, Petri WA Jr. Two cases of rarely recognized infection with Entamoeba moshkovskii Am J Trop Med Hyg. 2007 Apr;76(4):723-4.
  • 49. Ayed SB, Aoun K, Maamouri N, Abdallah RB, Bouratbine A. First molecular identification of Entamoeba moshkovskii in human stool samples in Tunisia. Am J Trop Med Hyg. 2008 Nov;79(5):706-7.
  • 50. Bouzid M, Kintz E, Hunter PR. Risk factors for Cryptosporidium infection in low and middle income countries: A systematic review and meta-analysis. PLoS Negl Trop Dis. 2018 Jun 7;12(6):e0006553.
  • 51. Feng Y, Ryan UM, Xiao L. Genetic Diversity and Population Structure of Cryptosporidium Trends Parasitol. 2018 Nov;34(11):997-1011.
  • 52. Khan A, Shaik JS, Grigg ME. Genomics and molecular epidemiology of Cryptosporidium species. Acta Trop. 2018 Aug;184:1-14.
  • 53. Xiao L. Molecular epidemiology of cryptosporidiosis: an update. Exp Parasitol. 2010 Jan;124(1):80-9.
  • 54. Zahedi A, Monis P, Gofton AW, Oskam CL, Ball A, Bath A, Bartkow M, Robertson I, Ryan U. Cryptosporidium species and subtypes in animals inhabiting drinking water catchments in three states across Australia. Water Res. 2018 May 1;134:327-40.
  • 55. Hamdy D, El-Badry A, Abd El Wahab W. Assessment of Giardia and Cryptosporidium Assemblages/Species and Their Viability in Potable Tap Water in Beni-Suef, Egypt Using Nested PCR/RFLP and Staining. Iran J Parasitol. 2019 Jul-Sep;14(3):368-78.
  • 56. Xiao L, Fayer R, Ryan U, Upton SJ. Cryptosporidium taxonomy: recent advances and implications for public health. Clin Microbiol Rev. 2004 Jan;17(1):72-97.
  • 57. Cacciò SM. Molecular epidemiology of human cryptosporidiosis. Parassitologia. 2005 Jun;47(2):185-92.
  • 58. Jiang J, Alderisio KA, Singh A, Xiao L. Development of procedures for direct extraction of Cryptosporidium DNA from water concentrates and for relief of PCR inhibitors. Appl Environ Microbiol. 2005 Mar;71(3):1135-41.
  • 59. Toledo J, Lombardelli J, Galarza R, Tiranti K, Garro CJ, Florin-Christensen M, Schnittger L, Tomazic ML. Comparison of DNA extraction methods to improve the molecular diagnosis of Cryptosporidium spp. from fecal samples of calves. Rev Inv Cient Univ Morón. 2018 Jul;2(4):17-28.
  • 60. Li N, Neumann NF, Ruecker N, Alderisio KA, Sturbaum GD, Villegas EN, 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 Sep 1;81(17):5845-54.
  • 61. Hadfield SJ, Robinson G, Elwin K, Chalmers RM. Detection and differentiation of Cryptosporidium spp. in human clinical samples by use of real-time PCR. J Clin Microbiol. 2011 Mar;49(3):918-24.
  • 62. Mary C, Chapey E, Dutoit E, Guyot K, Hasseine L, Jeddi F, Menotti J, Paraud C, Pomares C, Rabodonirina M, Rieux A, Derouin F; ANOFEL Cryptosporidium National Network. 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 Aug;51(8):2556-63.
  • 63. Rousseau A, La Carbona S, Dumètre A, Robertson LJ, Gargala G, Escotte-Binet S, Favennec L, Villena I, Gérard C, Aubert D. Assessing viability and infectivity of foodborne and waterborne stages (cysts/oocysts) of Giardia duodenalis, Cryptosporidium spp., and Toxoplasma gondii: a review of methods. Parasite. 2018;25:14.
  • 64. Hussein EM. Molecular identification of Cyclospora spp. using multiplex PCR from diarrheic children compared to others conventional methods. J Egypt Soc Parasitol. 2007 Aug;37(2):585-98.
  • 65. Li W, Kiulia NM, Mwenda JM, Nyachieo A, Taylor MB, Zhang X, Xiao L. Cyclospora papionis, Cryptosporidium hominis, and human-pathogenic Enterocytozoon bieneusi in captive baboons in Kenya. J Clin Microbiol. 2011 Dec;49(12):4326-9.
  • 66. Qvarnstrom Y, Benedict T, Marcet PL, Wiegand RE, Herwaldt BL, da Silva AJ. Molecular detection of Cyclospora cayetanensis in human stool specimens using UNEX-based DNA extraction and real-time PCR. Parasitology. 2018 Jun;145(7):865-870.
  • 67. Mathis GF, McDougald LR. Drug responsiveness of field isolates of chicken Coccidia. Poult Sci. 1982 Jan;61(1):38-45.
  • 68. Cruvinel LB, Nicaretta JE, Bastos TSA, Couto LFM, Santos JBD, Zapa DMB, Cavalcante ASA, Cruz BC, Borges DGL, Borges FA, Soares VE, Lopes WDZ. Eimeria species in dairy and beef cattle of different ages in Goiás state, Brazil. Rev Bras Parasitol Vet. 2018 Apr-Jun;27(2):169-76.
  • 69. Dubey JP, Bauer C. A review of Eimeria infections in horses and other equids. Vet Parasitol. 2018 May 30;256:58-70.
  • 70. Stensvold CR, Lewis HC, Hammerum AM, Porsbo LJ, Nielsen SS, Olsen KE, Arendrup MC, Nielsen HV, Mølbak K. Blastocystis: unravelling potential risk factors and clinical significance of a common but neglected parasite. Epidemiol Infect. 2009 Nov;137(11):1655-63.
  • 71. Noradilah SA, Moktar N, Anuar TS, Lee IL, Salleh FM, Manap SNAA, Mohtar NSHM, Azrul SM, Abdullah WO, Nordin A, Abdullah SR. Molecular epidemiology of blastocystosis in Malaysia: does seasonal variation play an important role in determining the distribution and risk factors of Blastocystis subtype infections in the Aboriginal community? Parasit Vectors. 2017 Jul 31;10(1):360.
  • 72. Alfellani MA, Stensvold CR, Vidal-Lapiedra A, Onuoha ES, Fagbenro-Beyioku AF, Clark CG. Variable geographic distribution of Blastocystis subtypes and its potential implications. Acta Trop. 2013 Apr;126(1):11-8.
  • 73. Tan KS, Mirza H, Teo JD, Wu B, Macary PA. Current Views on the Clinical Relevance of Blastocystis spp. Curr Infect Dis Rep. 2010 Jan;12(1):28-35.
  • 74. El Safadi D, Cian A, Nourrisson C, Pereira B, Morelle C, Bastien P, Bellanger AP, Botterel F, Candolfi E, Desoubeaux G, Lachaud L, Morio F, Pomares C, Rabodonirina M, Wawrzyniak I, Delbac F, Gantois N, Certad G, Delhaes L, Poirier P, Viscogliosi E. Prevalence, risk factors for infection and subtype distribution of the intestinal parasite Blastocystis sp. from a large-scale multi-center study in France. BMC Infect Dis. 2016 Aug 26;16(1):451.
  • 75. Tito RY, Chaffron S, Caenepeel C, Lima-Mendez G, Wang J, Vieira-Silva S, Falony G, Hildebrand F, Darzi Y, Rymenans L, Verspecht C, Bork P, Vermeire S, Joossens M, Raes J. Population-level analysis of Blastocystis subtype prevalence and variation in the human gut microbiota. Gut. 2019 Jul;68(7):1180-9.

  • Funding

    This work was funded by the University of Medical Sciences (UCIMED) of Costa Rica.
  • Ethics approval

    This study was approved by the Ethics Committee of the University of Medical Sciences, under project number CI-00218.

Publication Dates

  • Publication in this collection
    06 May 2022
  • Date of issue
    2022

History

  • Received
    01 Sept 2021
  • Accepted
    26 Jan 2022
Centro de Estudos de Venenos e Animais Peçonhentos (CEVAP/UNESP) Av. Universitária, 3780, Fazenda Lageado, Botucatu, SP, CEP 18610-034, Brasil, Tel.: +55 14 3880-7693 - Botucatu - SP - Brazil
E-mail: editorial.jvatitd@unesp.br