**About the Editor**

#### **David Gonz ´alez-Barrio**

David Gonzalez-Barrio is a graduate in Veterinary Medicine by the University of C ´ ordoba ´ (Spain) in 2009, and Master in wildlife health in 2012 by Spanish Wildlife Research Institute (IREC-CSIC-UCLM). Dr. Gonzalez-Barrio earned his European framework PhD in 2015 at the Spanish ´ Wildlife Research Institute (IREC) of the University of Castilla-La Mancha (UCLM) and Superior Council of Scientific Research (CSIC) in Spain after research training periods in Italy and Netherlands. During this time, he aimed to develop a PhD Thesis on epidemiology and molecular typing of *Coxiella burnetii* from domestic and wild animals in Spain. His research experience is based in animal health and zoonosis with multidisciplinary skills in the fields of animal disease diagnosis, disease epidemiology, molecular epidemiology and animal disease prevention and control, and its public health involvement in zoonotic diseases. Currently, he is a Researcher at the Toxoplasmosis and Intestinal Protozoa Unit of Parasitology Reference and Research Laboratory, Spanish National Centre for Microbiology, Health Institute Carlos III from Spain.

### *Editorial* **Zoonoses and Wildlife: One Health Approach**

**David González-Barrio**

Parasitology Reference and Research Laboratory, Spanish National Centre for Microbiology, Health Institute Carlos III, Ctra. Majadahonda-Pozuelo Km 2, Majadahonda, 28220 Madrid, Spain; dgonzalezbarrio@gmail.com or david.gonzalezb@isciii.es

Throughout history, wildlife has been an important source of infectious diseases transmissible to humans. Today, zoonoses with a wildlife reservoir constitute a major public health problem, affecting all continents. The importance of such zoonoses is increasingly recognized, and the need for more attention in this area is being addressed. The total number of zoonoses is unknown; some 1415 known human pathogens have been catalogued, and 62% are of zoonotic origin [1]. Over time, more and more human pathogens are found to be of animal origin. Moreover, most emerging infectious diseases in humans are zoonoses. Wild animals seem to be involved in the epidemiology of most zoonoses, and serve as major reservoirs for the transmission of zoonotic agents to domestic animals and humans [2]. The concept of the 'One Health' approach—involving collaboration between veterinary and medical scientists, policymakers, and public health officials—is necessary in order to foster joint cooperation and control of emerging zoonotic diseases [3]. Zoonotic diseases caused by a wide range of arthropods, bacteria, helminths, protozoans, and viruses can cause serious and even life-threatening clinical conditions in animals, with a number of them also affecting the human population due to their zoonotic potential.

The aim of the present Special Issue is to cover recent and novel research trends in zoonotic diseases in wildlife, including the relevant topics related to wildlife, zoonosis, public health, emerging diseases, infectious diseases, and parasitic diseases.

A total of 12 papers have been contributed by 96 authors from 14 countries to this issue, comprising 10 research articles, 1 communication, and 1 brief report (Figure 1). The number of specimens studied in this issue amounts to 5132, including wild animals, wild animals kept in captivity, domestic animals, and ticks; even human samples have been analyzed. More than 50 different species—including wild and domestic ungulates (e.g., red deer, roe deer, fallow deer, chamois, mouflon, European bison, wild boar, sheep, goat, cattle), wild carnivores (e.g., wolf, Eurasian lynx, Eurasian badger, coypu, beech marten, golden jackal), micromammals (e.g., yellow-necked field mouse, long-tailed field mouse, European water vole, white-toothed shrew, garden dormouse, common vole, house mouse, western Mediterranean mouse, black rat, Eurasian red squirrel), non-human primates (the genera *Cebuella, Cercocebus, Cercopithecus, Eulemur, Hylobates, Lemur, Macaca, Mandrillus, Saimiri, and Varecia*), turtles (e.g., *Testudo hermanni*, *T. h. boettgeri*, *T. graeca*, and *T. marginata*), bats (the families Pteropodidae, Emballonuridae, Rhinolophidae, Hipposideridae, and Vespertilionidae), and ticks (*Ixodes ricinus*, *Dermacentor marginatus*, *Hyalomma marginatum*) are included. Regarding the zoonotic pathogens represented in this issue, the presence of or exposure to 17 different pathogens—including viruses [4] (West Nile virus), bacteria [5–13] (*Anaplasma phagocytophilum, Coxiella burnetii, Helicobacter pylori, H. suis, Mycobacterium tuberculosis* Complex, *Salmonella* sp., and *Leptospira interrogans* sensu stricto), and parasitic protists [14,15] (e.g., *Cryptosporidium* spp., *Giardia duodenalis*, *Blastocystis* sp., *Enterocytozoon bieneusi*, *Entamoeba histolytica*, *Entamoeba dispar*, *Balantioides coli*, *Troglodytella* spp., *Leishmania* spp.)—are presented.

**Citation:** González-Barrio, D. Zoonoses and Wildlife: One Health Approach. *Animals* **2022**, *12*, 480. https://doi.org/10.3390/ ani12040480

Received: 25 January 2022 Accepted: 8 February 2022 Published: 15 February 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the author. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

**Figure 1.** A word cloud created from the titles of every article published in this Special Issue.

— — — − − The study of zoonotic pathogens present in wildlife mainly involves serological and/or molecular analyses, among others, for their detection, which is somewhat costly due to the difficulty in obtaining the necessary samples for analysis and ensuring that they are of high quality [16]; therefore, samples are often obtained from wild animals kept in captivity or in rescue centers [7,14]. In addition, the study of parasites involves searching for them, or their DNA—mainly in the feces of animals. In remote areas or resource-poor settings where the cold chain cannot be maintained, preservation and conservation of biological specimens including fecal samples—is a challenge; for this reason, Köster et al. [15] evaluated the suitability of filter cards for the long-term storage of fecal samples of animal and human origin that were positive for the diarrhea-causing protozoan parasites *Giardia duodenalis* and *Cryptosporidium hominis*. For this purpose, three commercially available Whatman ® filter cards were comparatively evaluated: the FTA® Classic card, the FTA® Elute Micro card, and the 903 Protein Saver card. *Giardia duodenalis* (*n* = 5)- and *C. hominis* (*n* = 5)-positive human stool samples were used to impregnate the selected cards at selected storage times (1 month, 3 months, and 6 months) and temperatures (−20 ◦C, 4 ◦C, and room temperature). Data presented by Köster et al. [15] demonstrate that Whatman ® cards are a cost-effective option for the preservation and long-term storage (up to six months) of fecal samples under a wide range of temperatures (from −20 ◦C to room temperature), without compromising their biospecimen stability and suitability for molecular-based diagnostic methods. Indeed, Whatman ® cards enable the molecular detection and genotyping of common diarrheacausing enteric protozoan parasites, including *C. hominis* and *G. duodenalis*.

— — A significant proportion of wildlife studies are carried out in conservation centers such as zoos—but also in wildlife rescue centers. Monitoring of infections that may be transmitted to humans by animals in wildlife rescue centers is very important in order to protect the staff engaged in rehabilitation practices. Casalino et al. [7] investigated the occurrence of non-typhoidal *Salmonella* in tortoises housed in a regional wildlife rescue

center in Apulia, Southern Italy, to assess the presence of *Salmonella* serovars that may pose a risk to operators involved in wildlife management. *Salmonella* may be a natural inhabitant of the intestinal tracts of turtles, rarely causing disease in turtles. This may represent a potential risk for humans, increasing the sanitary risk for operators in wildlife rescue centers. Casalino et al. [7] tested 69 adult turtles (*Testudo hermanni*, *T*. *h*. *boettgeri*, *T*. *graeca*, and *T*. *marginata*); the distribution of *Salmonella* spp. was significantly higher in *T*. *hermanni* than in other species. Two different *Salmonella* species (*S*. *enterica* and *S*. *bongori*) three *S*. *enterica* subspecies (*enterica*, *diarizonae*, and *salamae*), and five different serovars (Hermannswerder, Abony, Ferruch, Richmond, and Vancouver) within the group *S*. *enterica* subspecies *enterica* were identified. Most of the detected *Salmonella* types may represent a potential risk to public health. Reducing turtles' stress in order to minimize *Salmonella* shedding, as well as adopting correct animal husbandry procedures and hygiene techniques, may be useful to minimize the risk of transmission of *Salmonella* to humans. In particular, the adoption of gloves to manage turtles is a relevant preventive measure. Nevertheless, the greater measure of prevention is information and education on the potential sanitary risks of each professional figure involved in wildlife management.

On the other hand, little information is currently available on the epidemiology and zoonotic potential of parasitic and commensal protist species in captive non-human primates (NHPs). Köster et al. [14] investigated the occurrence, molecular diversity, and potential transmission dynamics of parasitic and commensal protist species in a zoological garden in southern Spain. The prevalence and genotypes of the main enteric protist species were investigated in fecal samples from NHPs (*n* = 51), zookeepers (*n* = 19), and free-living rats (*n* = 64) via molecular (PCR and sequencing) methods between 2018 and 2019. The presence of *Leishmania* spp. was also investigated in tissues from sympatric rats using PCR. *Blastocystis* sp. (45.1%), *Entamoeba dispar* (27.5%), *Giardia duodenalis* (21.6%), *Balantioides coli* (3.9%), and *Enterocytozoon bieneusi* (2.0%) (but not *Troglodytella* spp.) were detected in NHPs. *Giardia duodenalis* (10.5%) and *Blastocystis* sp. (10.5%) were identified in zookeepers, while *Cryptosporidium* spp. (45.3%), *G. duodenalis* (14.1%), and *Blastocystis* sp. (6.25%) (but not *Leishmania* spp.) were detected in rats. *Blastocystis* ST1, ST3, and ST8, along with *G. duodenalis* sub-assemblage AII, were identified in NHPs, and *Blastocystis* ST1 was identified in zookeepers. In rats, four *Cryptosporidium* (*C. muris*, *C. ratti*, and rat genotypes IV and V), one *G. duodenalis* (assemblage G), and three *Blastocystis* (ST4) genetic variants were detected. These results indicate high exposure of NHPs to zoonotic protist species. In conclusion, strong evidence of the occurrence of zoonotic *Blastocystis* transmission between NHPs and their handlers was provided, despite the use of personal protective equipment and the implementation of strict health and safety protocols. Free-living sympatric rats are infected by host-specific species/genotypes of the investigated protists, and seem to play a limited role as a source of infections to NHPs or humans in this setting.

Interactions taking place between sympatric wildlife/livestock/humans may contribute to interspecies transmission of pathogens [17]—this is the case of the *Mycobacterium tuberculosis* complex [18]. Mycobacteria can cause medically and socioeconomically significant diseases, including several non-tuberculous infections and tuberculosis, and are considered a One Health challenge due to their impact on public and animal health. These microorganisms are maintained and shared between the environment, domestic and wild animals, and humans. In this Special Issue, two studies are related to the interaction between domestic and wild species and the detection of mycobacteria in wild species such as badgers. Varela-Castro et al. [6] characterized the interactions that take place between several wild mammals and cattle via camera-trapping in order to provide insights into the dynamics of mycobacterial transmission opportunities in the environment of cattle farms located in Atlantic habitats in the northern Iberian Peninsula. Camera traps were set during a one-year period in cattle farms with a history of tuberculosis and/or non-tuberculous mycobacteriosis. A total of 1293 visits were recorded during 2741 days of camera observation. Only 23 visits showed direct contacts with cattle, suggesting that mycobacterial transmission at the wildlife–livestock interface occurs mainly through indirect interactions.

Results showed that cattle pastures represented the most appropriate habitat for interspecies transmission of mycobacteria, and badgers' latrines appear to be a potential hotspot for mycobacterial circulation between badgers, wild boars, foxes, and cattle. According to both previous epidemiological information and the interaction patterns observed, wild boars, badgers, foxes, and small rodents are the species or groups most often in contact with livestock and, thus, may be the most involved in the epidemiology of mycobacteriosis in the wildlife–livestock interface in this area. As Valera-Castaro et al. [6] pointed out in their work, the badger and its latrines are a hotspot for interspecies transmission—both domestic and wild; more specifically, Blanco Vázquez et al. [9] investigated the prevalence, spatial distribution, and temporal distribution of tuberculosis in 673 free-ranging Eurasian badgers (*Meles meles*) in Asturias (Atlantic Spain) between 2008 and 2020. The study's objective was to assess the role of badgers as a tuberculosis reservoir for cattle and other sympatric wild species in the region. Serum samples were tested in an in-house indirect P22 ELISA to detect antibodies against the *Mycobacterium tuberculosis* complex (MTC). In parallel, data on MTC isolation and single intradermal tuberculin test results were extracted for cattle that were tested and culled as part of the Spanish National Program for the Eradication of Bovine Tuberculosis. A total of 27/639 badgers (4.23%) were positive for MTC based on bacterial isolation, while 160/673 badgers (23.77%) were found to be positive with the P22 ELISA. The rate of seropositivity was higher among adult badgers than sub-adults. The authors found that the tuberculosis status of badgers in Asturias during 2008–2020 was associated with the tuberculosis status of local cattle herds, and results could not determine the direction of possible interspecies transmission, but they were consistent with the idea that the two hosts may exert infection pressure on one another. Both studies highlight the importance of monitoring this multi-host infection and disease in wildlife during epidemiological interventions in order to optimize outcomes under the One Health concept.

Deadly emerging and re-emerging zoonotic pathogens are transmitted mostly from wildlife reservoirs to humans or other animals during spillover events, with or without a vector intervention. In this special issue, two papers are included in which vectorborne zoonotic pathogens were studied. Ain-Najwa et al. [4] highlight the first evidence of West Nile virus (WNV) infection—a mosquito-borne virus—in Malaysian macaques and bats. Of the 81 macaques from mangrove forests sampled, 24 of the long-tailed macaques were seropositive for WNV, indicating that they were exposed to the virus; meanwhile, 5 out of 41 bats that were found in the caves from northern Peninsular Malaysia showed susceptibility to WNV. The authors found a high WNV antibody prevalence in macaques and a moderate WNV RNA in various Malaysian bat species, suggesting that WNV circulates through Malaysian wild animals, and that Malaysian bat species may be susceptible to the WNV infection. On the other hand, Grassi et al. [12] researched the genetic variants of *Anaplasma phagocytophilum* (a tick-borne pathogen causing zoonotic disease) in wild ungulates (the leading reservoir species) and feeding ticks (the main vector of infection) from northeastern Italy. Using biomolecular tools and phylogenetic analysis, ecotypes I and II were detected in both ticks (*Ixodes ricinus* species) and wild ungulates. Specifically, ecotype II was mainly detected in roe deer and related ticks, while ecotype I—the potentially zoonotic variant—was detected in *Ixodes ricinus* ticks, and also in wild ungulates. These findings reveal not only the wide diffusion of *Anaplasma phagocytophilum*, but also the presence of zoonotic variants.

Žele-Vengušt et al. [5] analyzed the exposure of free-ranging wild animals to zoonotic *Leptospira interrogans* sensu stricto in Slovenia; for this, blood samples from 249 wild animals between 2019 and 2020 were tested using the microscopic agglutination test for specific antibodies against the *Leptospira* serovars Icterohaemorrhagiae, Bratislava, Pomona, Grippotyphosa, Hardjo, Sejroe, Australis, Autumnalis, Canicola, Saxkoebing, and Tarassovi. Antibodies to at least one of the pathogenic serovars were detected in 77 (30.9%; CI = 25–37%) sera. The proportion of positive samples varied intraspecifically, and was the greatest in large carnivores (86%), followed by mesopredators (50%) and large herbivores

(17%). Out of the 77 positive samples, 42 samples (53.8%) had positive titers against a single serovar, while 35 (45.4%) samples had positive titers against two or more serovars. The most frequently detected antibodies were those against the serovar Icterohaemorrhagiae. This study confirmed the presence of multiple pathogenic serovars in wildlife throughout Slovenia. It can be concluded that wild animals are reservoirs for at least some of the leptospiral serovars, and are a potential source of leptospirosis for other wild and domestic animals, as well as for humans.

In their study, Cortez Nunes et al. [13] investigated the presence of *Helicobacter pylori* and *H. suis* DNA in free-range wild boars. *Helicobacter pylori* and *H. suis* are associated with gastric pathologies in humans. Interactions between domestic animals, wildlife, and humans can increase the risk of bacterial transmission between species. Samples of the gastric tissue of 14 free range wild boars (*Sus scrofa*) were evaluated for the presence of *H. pylori* and *H. suis* using PCR. Two samples were PCR-positive for *H. pylori*, and another for *H. suis*. These findings indicate that these microorganisms were able to colonize the stomachs of wild boars, and raise awareness of their putative intervention in the transmission cycle of *Helicobacter* spp..

Finally, this Special Issue includes three articles dealing with the potential role of livestock and wildlife as potential sources of human Q fever. Q fever is a worldwidedistributed zoonosis caused by *Coxiella burnetii*—a small intracellular bacterium belonging to γ-Proteobacteria that infects a wide range of animal species, including mammals, birds, and arthropods. People are infected through inhalation of aerosols contaminated with the bacteria expelled by infected animals during abortion or normal deliveries. Domestic ruminants, sheep, and goats are considered the main reservoirs of the infection and the principal source of human outbreaks. *Coxiella burnetii* has a complex ecology that replicates in multiple host species; however, the role of wildlife in its transmission is poorly understood. Krzysiak et al. [11] examined 523 serum samples obtained from European bison for the presence of specific antibodies in order to assess whether infection occurs in this species, and whether European bison may be an important source of infection in the natural environment, as suggested by historical reports. Only one (0.19%) serum sample was positive in ELISA, and two other samples were doubtful; the only seropositive animal was a free-living bull. This suggests possible transmission from domestic cattle by sharing pastures. The transmission of *C*. *burnetii* into the European bison was rather accidental in the country, and its role as an important wild reservoir is unlikely. In their study, González-Barrio et al. [10] examined spleen samples from 816 micromammals of 10 species, and 130 vaginal swabs from *Microtus arvalis* by qPCR, to detect *C. burnetii* infection and shedding, respectively; 9.7% of the spleen samples were qPCR-positive. The highest infection prevalence (10.8%) was found in *Microtus arvalis*, in which *C. burnetii* DNA was also detected in 1 of the 130 vaginal swabs (0.8%) analyzed. Positive samples were also found in *Apodemus sylvaticus* (8.7%), *Crocidura russula* (7.7%), and *Rattus rattus* (6.4%). Positive samples were genotyped by coupling PCR with reverse line blotting, and a genotype II+ strain was identified for the first time in one of the positive samples from *M. arvalis*, whereas only partial results could be obtained for the rest of the samples. Acute Q fever was diagnosed in one of the researchers who participated in the study, and it was presumably linked to *M. arvalis* handling. The results of the study are consistent with previous findings suggesting that micromammals can be infected by *C. burnetii*. The authors additionally suggest that micromammals may be potential sources to trace back the origin of human Q fever and animal coxiellosis cases in Europe, and might be relevant in the maintenance of wild-type *C. burnetii* strains that can be a matter of concern for animal and human health authorities. Espí et al. [8] investigated the seroprevalence of *C. burnetii* in domestic ruminants and wild ungulates, as well as the current situation of Q fever in humans, in a small region in northwestern Spain, where close contact at the wildlife–livestock–human interface exists, and information on *C. burnetii* infection is scarce. Seroprevalence of *C. burnetii* was 8.4% in sheep, 18.4% in cattle, and 24.4% in goats. Real-time PCR analysis of environmental samples collected in 25 livestock farms detected *Coxiella* DNA in dust and/or aerosols collected

in 20 of them. Analysis of sera from 327 wild ungulates revealed lower seroprevalence than that found in domestic ruminants. Exposure to the pathogen in humans was determined by IFAT analysis of 1312 blood samples collected from patients admitted to healthcare centers with Q-fever-compatible symptoms, such as fever and/or pneumonia. Results showed that 15.9% of the patients had IFAT titers ≥ 1/128, suggestive of probable acute infection. This study is an example of a One Health approach with medical and veterinary institutions involved in investigating zoonotic diseases.

Overall, the papers in this Special Issue reveal different perspectives of current research on zoonotic disease and wildlife, from applied field studies to investigations into the intricate mechanisms involved in the interaction between pathogens, wildlife, livestock, and humans.

**Funding:** This research received no external funding.

**Acknowledgments:** I would like to thank all of the authors who contributed their papers to this Special Issue, and the reviewers for their helpful recommendations. I am also grateful to all members of the *Animals* Editorial Office for giving me this opportunity, and for their continuous support in managing and organizing this Special Issue. Finally, I would like to acknowledge the academic editors Julie Arsenault, Fulvio Marsilio, Scott C. Williams, Nicole Gottdenker, Stefania Perrucci, and Laila Darwich Soliva, who made decisions for certain papers.

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**


### *Article* **Long-Term Preservation and Storage of Faecal Samples in Whatman ® Cards for PCR Detection and Genotyping of** *Giardia duodenalis* **and** *Cryptosporidium hominis*

**Pamela Carolina Köster 1 , Begoña Bailo 1 , Alejandro Dashti 1 , Carolina Hernández-Castro 1,2 , Rafael Calero-Bernal 3 , Francisco Ponce-Gordo 4 , David González-Barrio 1, \* and David Carmena 1, \***


**Simple Summary:** Preservation and storage of biological samples prior to testing and analysis is a pressing issue in the epidemiological field studies conducted in remote or poor-resource areas with limited or no access to electricity where the cold chain cannot be maintained. This is particularly true for faecal specimens of human and animal origin exposed to high degradation rates under environmental conditions characterised by high temperatures and humidity, such as those present in tropical and subtropical regions. Under this scenario, simple, safe, and cost-effective methods are highly needed to allow the collection and transportation of well-preserved faecal samples intended for pathogen detection without compromising the performance, reliability, and accuracy of molecular procedures methods used for detection and genotyping purposes. This study assessed the suitability of three commercially available filter cards for the preservation of faecal samples containing common diarrhoea-causing enteric protozoan parasites at different storage periods and temperature conditions. Obtained results demonstrated that filter cards impregnated with faecal matrices containing these pathogens are fully compatible with downstream molecular methods for up to six months at room temperature. Therefore, filter cards can be used for the safe transportation, preservation, and storage of faecal samples without the need of the cold chain.

**Abstract:** Preservation and conservation of biological specimens, including faecal samples, is a challenge in remote areas or poor-resource settings where the cold chain cannot be maintained. This study aims at evaluating the suitability of filter cards for long-term storage of faecal samples of animal and human origin positive to the diarrhoea-causing protozoan parasites, *Giardia duodenalis* and *Cryptosporidium hominis*. Three commercially available Whatman ® Filter Cards were comparatively assessed: the FTA® Classic Card, the FTA® Elute Micro Card, and the 903 Protein Saver Card. Human faecal samples positive to *G. duodenalis* (*n* = 5) and *C. hominis* (*n* = 5) were used to impregnate the selected cards at given storage (1 month, 3 months, and 6 months) periods and temperature (−20 ◦C, 4 ◦C, and room temperature) conditions. Parasite DNA was detected by PCR-based methods. Sensitivity assays and quality control procedures to assess suitability for genotyping purposes were conducted. Overall, all three Whatman ® cards were proven useful for the detection and molecular characterisation of *G. duodenalis* and *C. hominis* under the evaluated conditions. Whatman ® cards represent a simple, safe, and cost-effective option for the transportation, preservation, and storage of faecal samples without the need of the cold chain.

**Citation:** Köster, P.C.; Bailo, B.; Dashti, A.; Hernández-Castro, C.; Calero-Bernal, R.; Ponce-Gordo, F.; González-Barrio, D.; Carmena, D. Long-Term Preservation and Storage of Faecal Samples in Whatman ® Cards for PCR Detection and Genotyping of *Giardia duodenalis* and *Cryptosporidium hominis*. *Animals* **2021**, *11*, 1369. https://doi.org/ 10.3390/ani11051369

Academic Editor: Stefania Perrucci

Received: 29 March 2021 Accepted: 10 May 2021 Published: 12 May 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

**Keywords:** filter card; faeces; transportation; storage; preservation; *Giardia duodenalis*; *Cryptosporidium hominis*; PCR

#### **1. Introduction**

Biological samples including blood, saliva, stools, urine, tissue, and cells have become increasingly valuable sources of genetic material for downstream DNA and RNA testing. Because PCR-based methods are extremely sensitive to the quality and purity of the starting nucleic acid material [1], the appropriate preservation of biospecimens during procurement, transportation, and storage is essential to maximise the success of laboratory analyses [2]. This is a pressing issue in field epidemiological studies conducted in nonclinical or remote locations where resources are scarce or lacking, or when the cold chain cannot be guaranteed. In these poor-resource settings, simple, safe, and cost-effective methods are highly needed to allow the collection and transportation of intact biospecimens without detrimental effect on their biophysical properties and diagnostic utility.

The use of solid supports, such as filter cards for the collection and analysis of biospecimens, began in 1961, when Robert Guthrie developed what is now known as the Guthrie Test by collecting drops of blood on filter paper for the detection of phenylketonuria in new-borns [3]. Since then, filter cards have been developed and commercialised by different companies [4], being the Whatman® FTA® card technology (Cytiva, Marlborough, MA, USA) one of the most frequently used. FTA® cards are cotton-based, cellulose paper containing chemicals that lyse cells on contact, denature proteins, remove contaminants, and protect DNA from degradation (including UV radiation) by immobilising it onto the card's matrix [5]. FTA® cards allow the collection, preservation, and shipment of biospecimens for subsequent DNA and RNA analysis in a small space and at room temperature reducing transportation cost. Because chemically-treated FTA® cards inactivate the pathogenic agents in the samples they are carrying, they do not usually require import/export permits. Some of them (e.g., the 903 Protein Saver Card) have been approved by the US Food and Drug Administration (FDA) as class 2 devices [6]. Whatman® cards have been successfully used for the transportation and storage of a wide range of biospecimens including blood and serum [7], saliva [8], tissue [9], urine [10], sperm [11], mucus [12], and cerebral spinal fluid [13], among others.

Additionally, Whatman® cards have been proven useful for the diagnosis of pathogenic blood parasites, including canine microfilariae [14] and malaria-causing *Plasmodium* species [15]. However, very few studies have attempted to evaluate their efficacy for the detection of enteric pathogens in stool samples [16–18]. The protozoan enteroparasites, *Giardia duodenalis* (syn. *G. intestinalis* and *G. lamblia*) and *Cryptosporidium* spp., are two of the major contributors to the global burden of diarrhoeal illness both in humans [19,20] and livestock [21,22] globally. In poor-resource settings, more than 200 million human cases of symptomatic giardiosis are reported annually [23], whereas cryptosporidiosis (primarily by *C. hominis* and *C. parvum*) is the second leading cause of diarrhoea and deaths (after rotavirus) in children younger than five years of age [24]. Remarkably, production (cattle, sheep, goats, horses, donkeys, Bactrian camels) and free-living (non-human primates, among others) animal species have all been demonstrated to be competent hosts for *C. hominis* globally, confirming that this *Cryptosporidium* species is indeed zoonotic [25].

This study aims at comparing the performance of three types of Whatman® cards for the medium-/long-term preservation and storage (up to six months) of faecal material at different temperatures for downstream PCR detection of *G. duodenalis* and *Cryptosporidium hominis.*

#### **2. Materials and Methods**

#### *2.1. Selected Whatman® Cards*

Three different commercially available Whatman® cards with specific properties for sample collection, storage capacity, and costs were selected for comparative performance purposes: Whatman® Classic Cards, FTA® Elute Micro Cards, and 903 Protein Saver Cards (GE Healthcare Ltd., Cardiff, UK) (Table 1).


**Table 1.** Main features of the Whatman® cards used in the present comparative study.

Whatman® FTA® Classic Cards contains chemical denaturants and a free radical scavenger that have the ability to lyse cells on contact, denature proteins, and protect DNA from degradation. The extracted DNA remains tightly bound to the matrix while cell membranes and organelles are lysed and proteins and inhibitors are washed away [26]. FTA® Elute Micro Cards contains a chaotropic salt. Cells are lysed upon contact and proteins remain tightly bound while DNA is isolated from the matrix in a solution free of inhibitors with a simple water elution procedure [27]. Whatman® 903 Protein saver Card is an untreated cotton fibber-based matrix. It does not stabilise nor protect DNA from degradation.

#### *2.2. Stool Samples*

Five fresh, independent stool samples of human origin with a positive result for *G. duodenalis* by real-time PCR (qPCR, see Section 2.6) with cycle threshold (Ct) values ranging from 29.0 to 34.3 (median: 31.3; standard deviation: 2.0) and confirmed as assemblage B, sub-assemblage BIV at the *gdh* locus by Sanger sequencing were selected for this study. Five fresh, independent stool samples of human origin with a positive result for *C. hominis* by nested small subunit ribosomal RNA (*ssu* rRNA)-PCR (see Section 2.7) and confirmed as *C. hominis* (genotype IbA10G2) by Sanger sequencing were also included (Table S1). Human samples were chosen by mere convenience in terms of accessibility and quantity, but the faecal material from non-human animal sources is equally valid.

Initial diagnosis and subsequent genotyping of the *G. duodenalis*- and *C. hominis*positive stool samples were conducted at the Parasitology Reference and Research Laboratory of the Spanish National Centre for Microbiology (Majadahonda, Spain). After faecal sample homogenisation, 200 mg aliquots (*n* = 27, enough to cover all the experimental conditions considered in the study, see below) were weighed and stored at 4 ◦C in clean 1.5 mL Eppendorf tubes. Therefore, 270 stool sample aliquots were prepared.

#### *2.3. Impregnation of Whatman® Cards*

To standardise the experimental conditions of the study, total sample areas of the three Whatman*®* cards compared in the present study were normalised taking into consideration the sample area for each card specified in Table 1. A single sampling area of the FTA® Classic Card (enough for impregnating 200 mg of faecal material, (Figure 1a) equalled to four sampling areas of the FTA® Elute Micro Card (Figure 1b) and to three sampling areas of the 903 Protein Saver Card (Figure 1c). Under this premise, original Whatman® cards were cut and rearranged, as shown in Figure 1, to allow the coverage of three (1 month, 3 months, and 6 months) storage periods. This arrangement was used in triplicate to test three (−20 ◦C, 4 ◦C, and room temperature) storage conditions. Room temperature was considered that in the range of 15 to 25 ◦C.

**Figure 1.** Standardisation of sampling areas for each Whatman® card used in the present study to assess storage periods and conditions of impregnated stool samples (**a**) FTA® Classic Card; (**b**) FTA® Elute Micro Card; (**c**) 903 Protein Saver Card.

Normalised sampling units for each Whatman® card were impregnated with 200 mg of each aliquoted stool sample described above using cotton swabs embedded in phosphatebuffered saline (PBS) to soften the faecal material and facilitate the impregnation process. Impregnated Whatman® cards were allowed to dry at room temperature and stored in individual zip-lock plastic bags containing silica desiccant to keep moisture level low at the periods and storage conditions evaluated.

#### *2.4. Sensitivity Assay*

To estimate the minimum amount of *G. duodenalis* DNA detectable by qPCR in positive stool samples impregnated in Whatman*®* cards, one stool sample positive to this pathogen was selected. Serial-halved amounts (200 mg, 100 mg, 50 mg, 25 mg, 12.5 mg, and 6.25 mg) of faecal material were weighed and aliquoted in clean 1.5 mL Eppendorf tubes and subsequently used to impregnate single sampling areas of the Whatman*®* FTA*®* Classic Card. Impregnated cards were allowed to dry at room temperature and stored in individual zip-lock plastic bags containing silica desiccant for 1 month at room temperature. This experiment was not conducted with *Cryptosporidium*-positive samples because no semi-quantitative qPCR method was available in our laboratory for the detection of this pathogen.

#### *2.5. DNA Extraction and Purification*

Genomic DNA was extracted and purified from impregnated Whatman® cards at each storage period and condition described in Sections 2.3 and 2.4 using the QIAamp DNA Stool Mini Kit (QIAGEN®, Hilden, Germany), following the manufacturer's instructions with minor modifications. Briefly, whole-normalised sampling surfaces of each compared Whatman® card were cut into small pieces using a sterilised scissor and transferred into clean 2 mL Eppendorf tubes containing 1 mL of Inhibitex buffer. After incubation at 95 ◦C for 10 min, the tubes were thoroughly vortexed and centrifuged at 13,000 rpm for 3 min. Then, 350 µL of the obtained supernatants were transferred to clean 1.5 mL Eppendorf tubes and the rest of the procedure was completed using the QIAcube (QIAGEN®) automated DNA extraction system. Purified genomic DNA (200 µL) was stored at 4 ◦C until downstream PCR testing.

#### *2.6. Molecular Detection and Characterisation of Giardia duodenalis*

Detection of *G. duodenalis* DNA was achieved using a real-time PCR (qPCR) method targeting a 62-bp region of the *ssu* rRNA) gene of the parasite, as described elsewhere [28]. Amplification reactions (25 µL) contained 3 µL of template DNA, 12.5 pmol of primers Gd-80F and Gd-127R, 10 pmol of probe, and 12.5 µL TaqMan® Gene Expression Master Mix

(Applied Biosystems, California, CA, USA). Detection of parasitic DNA was performed on a Corbett Rotor GeneTM 6000 real-time PCR system (QIAGEN®). Water (no template) and genomic DNA (positive) controls were included in each PCR run.

For genotyping purposes, a semi-nested PCR was used to amplify a 432-bp fragment of the glutamate dehydrogenase (*gdh*) of *G. duodenalis* [29]. Briefly, PCR reaction mixtures (25 µL) included 5 µL of template DNA and 0.5 µM of the primer pairs GDHeF/GDHiR in the primary reaction and GDHiF/GDHiR in the secondary reaction.

#### *2.7. Molecular Detection and Characterisation of Cryptosporidium hominis*

Detection of *C. hominis* DNA was achieved using a nested-PCR protocol to amplify a 587-bp fragment of the *ssu* rRNA gene of the parasite as described elsewhere [30]. Amplification reactions (50 µL) included 3 µL of DNA sample and 0.3 µM of the primer pairs CR-P1/CR-P2 in the primary reaction and CR-P3/CPB-DIAGR in the secondary reaction. Reaction mixes also contained 2.5 units of MyTAQ™ DNA polymerase (Bioline GmbH, Luckenwalde, Germany) and 5× MyTAQ™ Reaction Buffer containing 5 mM dNTPs and 15 mM MgCl2.

For genotyping purposes, a nested PCR was used to amplify an 870-bp fragment of the 60 kDa glycoprotein (*gp60*) of *C. hominis* [31]. PCR reaction mixtures (50 µL) included 2-3 µL of template DNA and 0.3 µM of the primer pairs AL-3531/AL-3535 in the primary reaction and AL-3532/AL-3534 in the secondary reaction.

The semi-nested and nested PCR protocols described above were conducted on a 2720 Thermal Cycler (Applied Biosystems). Reaction mixes included 2.5 units of MyTAQ™ DNA polymerase (Bioline GmbH, Luckenwalde, Germany), and 5× MyTAQ™ Reaction Buffer containing 5 mM dNTPs and 15 mM MgCl2. Amplicons were visualised under UV light after 2% agarose gel electrophoresis. Positive-PCR products were directly sequenced in both directions using inner primer sets. DNA sequencing was conducted by capillary electrophoresis using the BigDye® Terminator chemistry (Applied Biosystems) on an on ABI PRISM 3130 automated DNA sequencer.

#### *2.8. Quality Control*

To confirm the suitability of the three compared Whatman® cards for genotyping purposes, purified genomic DNAs from single *G. duodenalis*- and *C. hominis*-positive samples stored for six months (the maximum period covered in the present study) at 4 ◦C and room temperature (the most sensitive conditions to DNA damage evaluated here) were re-amplified by *gdh*-PCR (*G. duodenalis*) and *gp60*-PCR (*C. hominis*) and sequenced as described above. The quality of the obtained chromatograms was visually inspected, and the accuracy of the readings confirmed by alignment with appropriate reference sequences retrieved from GenBank.

#### *2.9. Statistical Analyses*

The Shapiro–Wilk's test was used to assess the normality of distribution of the Ct values obtained in *G. duodenalis*-positive samples during qPCR analyses at each period and storage condition evaluated. Once normality was demonstrated, an analysis of variance (ANOVA) for simultaneous comparison of conditions was conducted. A probability (*p*) value < 0.05 was considered evidence of statistical significance. Statistical analyses were performed using the R-software version 4.0.2 [32].

#### **3. Results**

#### *3.1. Performance of Whatman® Cards for the Preservation and Storage of G. duodenalis-Positive Faecal Samples*

All except two samples (an FTA*®* Elute Micro Card stored at room temperature for 1 month, and an FTA*®* Classic Card stored at -20 ◦C for three months) tested positive for *G. duodenalis* by qPCR, yielding Ct values similar to those generated at the time of initial diagnosis (Table S1). At 1 month-length storage, all three Whatman*®* cards performed

equally well, with those kept at 4 ◦C yielding lower (but not statistically significant, *p* = 0.96) Ct values (Figure 2a). At 3 months-length storage, the FTA*®* Classic Card provided the best diagnostic values in terms of sensitivity (lower Ct values) and precision (lower standard deviation) regardless of the temperature. These results were statistically significant when compared with those obtained with the 903 Protein Saver Card (*p* = 0.01), but not with the FTA*®* Elute Micro Card (*p* = 0.40) (Figure 2b). No statistically significant differences (*p* = 0.99) were observed among the three compared Whatman*®* cards at 6 months-length storage (Figure 2c). It should be noted that the difference observed between the FTA*®* Classic Card and the 903 Protein Saver Card in Figure 2b is associated to the effect caused by an outlier data value generated with the latter at 4 ◦C storage conditions. Removal of this value resulted in the loss of statistical significance.

**Figure 2.** *Cont*.

**Figure 2.** Box plot of cycle threshold (Ct) values generated from *Giardia duodenalis* isolates extracted from the three Whatman® cards evaluated in the present study at different storage conditions. (**a**): 1 month-length storage; (**b**): 3 months-length storage; (**c**): 6 month-length storage. Horizontal thick lines within boxes represent median values Upper and lower whiskers represent the data range. Plotted dots represent outliers. Using the Tukey's Honestly Significant Difference test as multiple post hoc comparison method, statistical significance is represented as \* (*p* < 0.05).

#### *3.2. Performance of Whatman® Cards for the Preservation and Storage of C. hominis-Positive Faecal Samples*

The generated *ssu*-PCR results after the processing and testing of *C. hominis*-positive stool samples impregnated in the Whatman*®* cards at the storage periods and conditions assessed in the present study are shown in Figure 3. All tested samples stored for 1 month (Figure 3a), 3 months (Figure 3b), or 6 months (Figure 3c) yielded clear amplicons regardless of the storage temperature considered (Figure S1).

**Figure 3.** Agarose gel electrophoresis (2% *w*/*v*) detection of PCR products showing the presence of a 587-bp amplicon for the *Cryptosporidium hominis ssu* rRNA gene extracted from the three Whatman® cards evaluated in the present study at different storage conditions. (**a**): 1 month-length storage; (**b**): 3 months-length storage; (**c**): 6 month-length storage.

#### *3.3. Sensitivity Assay*

When tested by qPCR, serial-halved amounts of faecal material containing *G. duodenalis* cysts impregnated in Whatman® FTA® Classic Cards generated Ct values ranging from 25.6 (corresponding to 200 mg of faeces) to 33.1 (corresponding to 6.3 mg of faeces) (Table 2).

**Table 2.** Cycle threshold (Ct) values obtained by real-time PCR in serially-halved amounts of a faecal sample positive for *G. duodenalis* impregnated in Whatman® FTA® Classic Cards.


#### *3.4. Quality Control*

Figure S1 shows the *gdh*-PCR and *gp60*-PCR amplification results obtained with the purified genomic DNAs from the two samples positive to *G. duodenalis* and *C. hominis*, respectively. Both samples were stored in all three compared Whatman® cards for 6 months (the maximum period covered in the present study) at 4 ◦C and room temperature (the most likely conditions to induce DNA damage evaluated here). Two of the six *G. duodenalis*-positive extracts were successfully amplified at the *gdh* locus (Figure S1a) and their associated chromatograms displayed good quality sequences confirming the identity (subassemblage BIV) of the parasite (Figure S2a). All *C. hominis*-positive extracts yielded clear amplicons at the *gp60* locus, irrespectively of the Whatman® card used or the temperature considered (Figure S1b). Sanger sequencing analysis revealed good quality sequence data confirming the identity (genotype IbA10G2) of the parasite (Figure S2b).

#### **4. Discussion**

This study evaluated the suitability of three commercially available Whatman® Filter Cards (the FTA® Classic Card, the FTA® Elute Micro Card, and the 903 Protein Saver Card) for the long-term storage of faecal material containing *G. duodenalis* cysts and *C. hominis* oocysts, two of the major contributors to the global burden of diarrhoeal illness both in humans [19,20] and livestock [21,22] globally. Of note, both protozoan parasites present aggregated distributions depending on the host species, genetic variants, or even geographical area considered [33–37]. Because both *G. duodenalis* and *Cryptosporidium* spp. are common findings in the faecal material of human and animal hosts, some of their species/genotypes have zoonotic potential, and are ubiquitous in the environment, research on the epidemiology and transmission of these pathogens should be always conducted under the One Health umbrella. This approach is particularly useful in those epidemiological scenarios where different epidemiological (e.g., domestic and sylvatic) cycles of the parasites overlap, allowing the occurrence of spillover events [33,34,36]. The three major contributions of this survey include the demonstration that (i) the three compared Whatman© cards performed near equally well in maintaining the stability of the faecal material for up to six months irrespectively of the storage temperature; (ii) the parasitic DNA extracted from impregnated Whatman® cards was suitable for subsequent molecular detection and genotyping purposes; and (iii) Whatman® cards represent simple, time- and cost-effective options for the safe storage and transportation of faecal samples of human and animal origin without the need of the cold chain.

FTA® card technology was originally designed as a matrix for blood storage and processing medium [5]. Because of their versatility and simplicity of use, FTA® cards were soon after tested for storing other biological samples including saliva [8], tissue [9], urine [10], sperm [11], mucus [12], and cerebral spinal fluid [13]. This tool has been

also assessed for the molecular detection of gastrointestinal parasites (e.g., the coccidian *Cryptosporidium* spp. and *Cyclospora cayetanensis*, the flagellated *G. duodenalis*, and the microsporidia *Encephalitozoon intestinalis*) in matrices including clinical specimens and fresh produce [16–18]. In a seminal study, FTA® card templates prepared from purified *Cryptosporidium* spp. oocysts and *E. intestinalis* spores and subsequently assessed by PCR allowed the identification of as few as 10 oocysts/spores. Similar results were also observed with clinical samples including faeces, urine, sputum, and foods (berries) [16]. The authors concluded that PCR analysis using the FTA® card format for DNA template preparation was routinely unaffected by the matrix from which the sample was derived while still maintaining a high level of detection sensitivity [16]. In a subsequent survey, known concentrations of *G. duodenalis* cysts and *Cryptosporidium* spp. oocysts were serially diluted, spiked into a faecal suspension from a pathogen-negative stool, and smeared onto FTA® Elute Micro cards [17]. Stool cards were then stored at room temperature, DNA was extracted and purified using QIAGEN protocols and tested by multiplex PCR coupled with Luminex assay at 1 week, 1 month, and 3 months. A limited number of stool cards were also stored at 4 ◦C and in a humid incubator at 31 ◦C for 1 week to determine the impact of environmental conditions on detection. The authors detected *G. duodenalis* at 3 months with a 2-log reduction from the original concentration, whereas *Cryptosporidium* spp. was undetected after 1 month of storage. Failure to detect the presence of *Cryptosporidium* spp. for longer periods of time was attributed to suboptimal breakage of the parasite oocyst wall [17]. Finally, the 903 Protein Saver card has been evaluated for the detection of *G. duodenalis*, *Cryptosporidium* spp., and *Entamoeba histolytica* in either whole faecal samples or stool suspensions using QIAGEN procedures for DNA purification and qPCR for detection [18]. In this study the cards were stored for only 48 h before DNA purification and qPCR testing. Depending on the starting (whole or suspension) faecal material used for impregnation, obtained overall sensitivities were 85–95% for *G. duodenalis*, 60–85% for *E. histolytica*, and 35–40% for *Cryptosporidium* spp. In general, faecal suspensions yielded poorest qPCR amplification results than whole faecal samples. Parasite load was identified as a critical factor for qPCR success [18].

This study improves current knowledge on the practicality and performance of Whatman® cards for the molecular detection of diarrhoea-causing enteric protozoan parasites in several aspects. First, this is (to author´s knowledge) the first attempt conducted to date to compare simultaneously three different types of Whatman® cards including the FTA® Classic Card, the FTA® Elute Micro Card, and the 903 Protein Saver Card. Previous studies focused on a specific card type only [16–18]. Second, the evaluated storage period has been extended to 6 months, three more months that the maximum period covered in previous studies [17]. Third, this survey evaluated the effect of three different storage temperatures. Freezing (–20 ◦C) and refrigeration (4 ◦C) temperatures represented the most common conditions in routine laboratory practice, whereas the room temperature condition attempted to mimic those present in field work characterised by lack of electric supply where sample conservation is a pressing issue. Previous studies were conducted primarily at room temperature [16–18], with only few impregnated cards being tested at other temperatures [17]. Fourth, present results were obtained exclusively with true clinical faecal samples, whereas those from previous studies were mostly derived from purified parasitic material [16] or artificially spiked stools [17]. Fifth, quality control data presented here provided evidence demonstrating that Whatman® cards were suitable for genotyping (in addition to detection) purposes, including Sanger sequencing. None of the studies carried out before assessed this possibility.

A major contribution of this study was the finding that all three compared Whatman® cards yielded sensitivity values near 100%, irrespectively of the storage period, the temperature considered, or the parasite species investigated. These figures were considerably higher than those reported in similar surveys [16–18]. Several factors may account, at least partially, for the differences observed. For instance, we used high impregnation loads (200 mg) of faecal material, in line with the recommendations of the QIAGEN procedure

used for DNA extraction and purification. In addition, our protocol included a modification (sample incubation with Inhibitex buffer at 95 ◦C for 10 min) specifically intended at improving the efficiency of cyst/oocyst breakage, an issue previously identified as a factor limiting the diagnostic sensitivity of PCR assays [17]. Variations in the diagnostic performance of the PCR methods used (conventional nested PCR, qPCR, multiplex qPCR) may also influence the amplification success rate obtained in these studies.

This study presents, however, some limitations that should be taken into consideration. For instance, a straightforward application of the data presented here is the potential usefulness of Whatman® cards as a convenient stool storage system for periods longer than 6 months (e.g., in biobanks), a possibility that should be conveniently evaluated in further studies. Of note, room temperature was considered here those in the range of 15 to 25 ◦C. More extreme temperature (and humidity) conditions, such as those typically present in tropical and sub-tropical regions, may affect the performance of the Whatman® cards. This possibility should be conveniently evaluated in future studies. Also, only faecal samples positive for *G. duodenalis* and *C. hominis* were investigated. Although we do not anticipate significant performance differences with other enteric protist species, this is also an issue that remains to be fully elucidated. Finally, our data can be used as proof of concept for the suitability of Whatman® technology for the safe storage and transportation of faecal material at room temperature without detrimental effects on stability and diagnostic features, although this fact should be demonstrated in *ad*-*hoc* studies.

#### **5. Conclusions**

Data presented here demonstrate that Whatman® cards are a cost-effective option for the preservation and long-term storage (up to six months) of faecal samples under a wide range of temperatures (from –20 ◦C to room temperature) without compromising their biospecimen stability and suitability for molecular-based diagnostic methods. Indeed, Whatman® cards enable the molecular detection and genotyping of common diarrhoeacausing enteric protozoan parasites, including *C. hominis* and *G. duodenalis*. Further research should be conducted to unambiguously demonstrate the usefulness of Whatman® cards in field epidemiological surveys involving larger number of faecal samples, wider ranges of temperature and humidity conditions, and storage periods longer than six months. In practical terms, Whatman® cards would allow the obtaining and safe transportation of faecal samples of human and animal origin from remote areas to clinical or research laboratories without the need of the cold chain.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/ani11051369/s1, Table S1: Real-time PCR cycle threshold values obtained after the amplification of *Giardia duodenalis*-positive stool samples impregnated in the Whatman® Cards at the storage periods and conditions assessed in the present study, Figure S1: Agarose gel electrophoresis (2% *w*/*v*) detection of PCR products used for evaluating the suitability of Whatman® cards for genotyping and Sanger sequencing purposes. (a) Results showing the presence of a 432-bp amplicon for the *Giardia duodenalis gdh* gene in sample G145. Some lanes have been cut and re-arranged to keep the same order in the whole figure; (b): Results showing the presence of an 870-bp amplicon for the *Cryptosporidium hominis gp60* gene in sample C578., Figure S2: Representative chromatograms showing Sanger sequencing results for PCR amplicons generated from genomic DNA extracted and purified from Whatman® Cards. (a): Results for the *Giardia duodenalis gdh* gene in sample G145; (b): Results for the *Cryptosporidium hominis gp60* gene in sample C578.

**Author Contributions:** Conceptualisation, P.C.K., R.C.-B., F.P.-G., and D.C.; methodology, P.C.K., B.B.; A.D., and C.H.-C.; validation, R.C.-B., F.P.-G., D.G.-B., and D.C.; formal analysis, P.C.K., A.D., R.C.-B., F.P.-G., D.G.-B., and D.C.; investigation, P.C.K.; resources, D.C.; writing—original draft preparation, P.C.K. and D.C.; writing—review and editing, P.C.K., R.C.-B., F.P.-G., D.G.-B., and D.C.; supervision, R.C.-B., F.P.-G., D.G.-B., and D.C.; project administration, D.C.; funding acquisition, D.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Health Institute Carlos III (ISCIII), Ministry of Science, Innovation and Universities (Spain), grant number PI16CIII/00024.

**Institutional Review Board Statement:** The study was conducted according to the guidelines of the Declaration of Helsinki and approved by the Ethics Committee of the Health Institute Carlos III (CEI PI17\_2017-v3; date of approval: 23 October 2017).

**Data Availability Statement:** All relevant data are within the article and its additional files.

**Acknowledgments:** David González-Barrio was recipient of a "Sara Borrell" postdoctoral fellow-ship (CD19CIII/00011) funded by the Spanish Ministry of Science, Innovation and Universities.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

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