A Review of Strongyloides spp. Environmental Sources Worldwide.

Strongyloides spp. are parasitic nematodes that are transmitted through the environment and are capable of causing disease. These nematodes affect an estimated 3-300 million humans worldwide. Identifying the environmental reservoirs of Strongyloides spp. is essential for the development of appropriate control strategies. This systematic literature review examined all published studies that identified Strongyloides stercoralis, Strongyloides fuelleborni, Strongyloides fuelleborni kellyi, and Strongyloides spp. from an environmental source. Most studies detected the nematode from dog and primate fecal samples. Other environmental sources identified were ruminants, cats, rodents, insects, water, soil, as well as fruit and vegetables. Most studies used microscopy-based identification techniques; however, several employed molecular-based techniques, which have become increasingly popular for the detection of Strongyloides spp. A limitation identified was a lack of studies that comprehensively screened all potential environmental samples in a region. Future research should undertake this holistic screening process to identify which environmental reservoirs pose the greatest significance to human health. Potential controls can be identified through the identification of environmental sources. Understanding where Strongyloides spp. is commonly found within the environment of endemic areas will inform environmental control strategies to reduce this neglected disease.

1. Introduction

Strongyloidiasis is a disease caused by parasitic nematodes of the genus Strongyloides. Within this genus, three species, Strongyloides stercoralis, Strongyloides fuelleborni, and Strongyloides fuelleborni kellyi are known to parasitize humans [1,2].

S. stercoralis, S. fuelleborni, and S. fuelleborni kellyi are capable of autoinfecting the host. This occurs after adult female parthenogenic nematodes within the infected human shed eggs. These eggs develop to larvae that are passed within the stool. A certain number burrow through the wall of the large intestine, thereby reinfecting the body. Infected individuals can have a low-level undetected infection for many years [3]. When this auto-infective life cycle becomes uncontrolled in immunocompromised, young, and elderly patients, a disseminated infection can develop. Disseminated infection occurs when the parasite travels throughout the body. This can result in sepsis, bacterial meningitis, or gastrointestinal hemorrhage [4]. The mortality rate from a disseminated infection and its comorbidities is estimated to be 80% [3]. The larvae that are passed within the stool are then capable of completing a free-living cycle, in which they molt twice to develop into filariform larvae. These infective filariform larvae are capable of then reinfecting humans, where they can be involved with the autoinfection cycle again [5].

Both S. stercoralis and S. fuelleborni are able to complete their life cycle within animals such as canids, primates, and insects. Animal species-specific strains of S. stercoralis unable to infect humans have been identified [6]. This ability for the nematode to reproduce within other animals indicates that all infected animals’ feces may pose an infection threat to humans.

After excretion in the stool, larvae can survive and reproduce within the environment, and environmental sources contaminated with larvae can cause reinfection. Although Strongyloides spp. are classified as soil-transmitted helminths, locations that harbor Strongyloides spp. within the environment, with the exception of soil, have not been investigated holistically [7,8]. By reviewing and collating all reported environmental sources of S. stercoralis, S. fuelleborni, and S. fuelleborni kellyi, environmental interventions can be implemented.

We need a better understanding of the environmental sources of Strongyloides spp.; resistance to the current anthelminthic drugs has been observed in other Strongyloides spp. [9]. Both environmental and clinical control of Strongyloides spp. is essential [10]. The aim of this review is to identify all research reporting S. stercoralis, S. fuelleborni, S. fuelleborni kellyi, and Strongyloides spp. within environmental sources worldwide.

2. Results

One thousand two hundred and twenty-two papers were retrieved from SCOPUS and Web of Science using the search terms identified as suitable, as seen in Table 1, with 174 articles identified as eligible for inclusion.

Table 1

Complete search strategy and all key words used to identify relevant literature.

Search Terms Employed to Identify Relevant Literature
Strongyloides OR Strongyloidiasis OR “Strongyloides stercoralis” OR “S. stercoralis” OR “Strongyloides fuelleborni” OR “S. fulleborni” OR “Strongyloides fulleborni kellyi” OR “S. fulleborni kellyi”
AND
“Tap Water” OR “Potable water” OR Water OR Soil OR Dirt OR sediment OR synanthropic OR “synanthropic insect” OR Insect OR “Musca domestica” OR flies OR “Musca vetustissima” OR Sarcophagidae OR “Chrysomya megacephala” OR “Musca sorbens” OR “Lucilia cuprina” OR “Calliphora vicina” OR “Blattella germanica” OR “Periplaneta Americana” OR Cockroach OR dog OR “Canis lupis” OR zoonotic OR Monkey OR “septic tank” OR waste OR wastewater OR rubbish OR trash OR environment

S. stercoralis was identified in 35% of all studies and S. fuelleborni in 10% of all studies; both S. stercoralis and S. fuelleborni were identified in 1% of all studies. S. fuelleborni and Strongyloides spp. were identified in 0.5% of studies, and genus-level identification was identififed in 55% of all studies, as seen in Table A1. S. fuelleborni kellyi was not identified within any papers.

Table A1

Summary of all reports, and studies identifying S. stercoralis, S. fuelleborni, S. fuelleborni kellyi, and Strongyloides spp. within environmental sources worldwide.

Species Parasite	Prevalence	Sample Size	Detection Method	Country	Reference	Source
Strongyloides stercoralis	1%	3465	Microscopy	Romania	(Ardelean et al., 2005) [60]	Dog
Strongyloides stercoralis	49%	35	Molecular	Australia	(Beknazarova et al., 2017) [59]	Dog
Strongyloides stercoralis	<1%	3208	Microscopy	Iceland	(Eydal and Skirnisson 2016) [88]	Dog
Strongyloides stercoralis	<1%	215	Microscopy	Brazil	(Ferreira et al., 2006) [89]	Dog
Strongyloides spp.	<1%	457	Microscopy	Canada	(Gaunt and Carr 2011) [90]	Dog
Strongyloides stercoralis	<1%	181	Microscopy	Brazil	(Goncalves et al., 2007) [91]	Dog
Strongyloides stercoralis	87%	88	Molecular	Cambodia	(Jaleta et al., 2017) [6]	Dog
Strongyloides stercoralis	<1%	879	Microscopy	Greece	(Kostopoulou et al., 2017) [48]	Dog
Strongyloides stercoralis	<1%	189	Microscopy	Thailand	(Leelayoova et al., 2009) [49]	Dog
Strongyloides stercoralis	45%	171	Microscopy	Brazil	(Martins et al., 2012) [58]	Dog
Strongyloides stercoralis	4%	52	Microscopy	Romania	(Mircean et al., 2012) [92]	Dog
Strongyloides spp.	5%	175	Microscopy	Malaysia	(Noor Azian et al., 2008) [40]	Dog
Strongyloides stercoralis	2%	281	Microscopy	Greece	(Papazahariadouet al., 2007) [50]	Dog
Strongyloides stercoralis	2%	272	Microscopy	Italy	(Paradies et al., 2017) [93]	Dog
Strongyloides spp.	11%	90	Microscopy	Sri Lanka	(Perera et al., 2013) [57]	Dog
Strongyloides stercoralis	6%	174	Microscopy	Iran	(Razmi et al., 2009) [52]	Dog
Strongyloides stercoralis	<1%	239	Microscopy	Italy	(Riggio et al., 2013) [53]	Dog
Strongyloides stercoralis	<1%	639	Microscopy	Italy	(Sauda et al., 2018) [55]	Dog
Strongyloides spp.	15%	94	Microscopy	Cambodia	(Schär et al.,2014) [87]	Dog
Strongyloides stercoralis	10%	60	Microscopy	Slovakia	(Štrkolcová et al., 2017) [45]	Dog
Strongyloides spp.	2%	171	Microscopy	England	(Wright et al., 2016) [56]	Dog
Strongyloides stercoralis	<1%	463	Microscopy	Italy	(Zanzani et al., 2014) [94]	Dog
Strongyloides spp.	2%	197	Microscopy	Thailand	(Pumidonming et al., 2016) [51]	Dog
Strongyloides stercoralis	18%	824	Microscopy	Qatar	(Abu-Madi et al., 2007) [95]	Cat
Strongyloides spp.	47%	28	Microscopy	Christmas Island	(Adams et al., 2008) [96]	Cat
Strongyloides spp.	54%	37	Microscopy	Brazil	(Lima et al., 2017) [97]	Cat
Strongyloides spp.	3%	414	Microscopy	Romania	(Mircean et al., 2010) [98]	Cat
Strongyloides stercoralis	14%	173	Microscopy	Brazil	(Monteiro et al., 2016) [99]	Cat
Strongyloides stercoralis	44%	103	Microscopy	Kenya	(Njuguna et al., 2017) [100]	Cat
Strongyloides spp.	<1%	300	Microscopy	Thailand	(Rojekittikhun et al., 2014) [101]	Cat
Strongyloides stercoralis	21%	38	Microscopy	Thailand	(Sedionoto and Anamnart 2018) [102]	Cat
Strongyloides spp.	99 1.0%	99	Microscopy	Denmark	(Takeuchi-Storm et al., 2015) [103]	Cat
Strongyloides fuelleborni	UNK	UNK	Molecular and Microscopy	Japan	(Arizono et al., 2012) [13]	Primate
Strongyloides spp.	41 44%	41	Microscopy	Uganda	(Bezjian et al., 2008) [11]	Primate
Strongyloides spp.	37%	24	Microscopy	French Guiana	(De Thoisy et al., 2001) [14]	Primate
Strongyloides spp.	21%	125	Microscopy	India	(Ekanayake et al., 2006) [104]	Primate
Strongyloides fuelleborni	28%	293	Microscopy	Uganda	(Gillespie et al., 2004) [105]	Primate
Strongyloides fuelleborni and Strongyloides stercoralis	<1% S. stercoralis, 4% S. fuelleborni	2103	Microscopy	Uganda	(Gillespie et al., 2005) [106]	Primate
Strongyloides fuelleborni	84%	153	Microscopy	Tanzania	(Gillespie et al., 2010) [107]	Primate
Strongyloides fuelleborni and Strongyloides spp.	11% S. fulleborni, 15% S. spp	27	Microscopy	Spain	(Gomez et al., 1996) [15]	Primate
Strongyloides fuelleborni	23%	401	Microscopy	Japan	(Gotoh 2000) [108]	Primate
Strongyloides fuelleborni and Strongyloides stercoralis	100%	7	Molecular	Uganda	(Hasegawa et al., 2016) [61]	Primate
Strongyloides spp.	88%	96	Microscopy	Ecuador	(Helenbrook et al., 2015) [64]	Primate
Strongyloides spp.	4%	238	Microscopy	Uganda	(Hodder and Chapman 2012) [109]	Primate
Strongyloides spp.	7%	40	Microscopy	Kenya	(Karere and Munene 2002) [12]	Primate
Strongyloides spp.	41%	624	Microscopy	Borneo	(Klaus et al., 2018) [62]	Primate
Strongyloides fuelleborni	32%	652	Microscopy	Borneo	(Klaus et al., 2017) [110]	Primate
Strongyloides fuelleborni	57%	141	Microscopy	Puerto Rico	(Knezevich et al., 1998) [111]	Primate
Strongyloides spp.	43%	686	Microscopy	Tanzania	(Kooriyama et al., 2012) [112]	Primate
Strongyloides spp.	74%	3142	Microscopy	Côte d’Ivoire	(Kouassi et al., 2015 [113]	Primate
Strongyloides spp.	13%	366	Microscopy	India	(Kumar et al., 2018) [114]	Primate
Strongyloides fuelleborni	44% S. fuelleborni, 4% S. spp.	25	Microscopy	Malaysia	(Kuze et al., 2010) [115]	Primate
Strongyloides fuelleborni	95%	20	Molecular and Microscopy	Indonesia	(Labes et al., 2011) [116]	Primate
Strongyloides spp.	37%	59	Microscopy	Ethiopia	(Legesse and Erko 2004) [117]	Primate
Strongyloides spp.	5%	222	Microscopy	Belgium	(Levecke et al., 2007) [118]	Primate
Strongyloides spp.	6%	3349	microscopy	China	(Li et al., 2017) [65]	Primate
Strongyloides stercoralis	6%	46	Microscopy	Nigeria	(Mafuyai et al., 2013) [119]	Primate
Strongyloides spp.	50%	134	Microscopy	Costa Rica	(Maldonado-Lopez et al., 2014) [120]	Primate
Strongyloides spp.	77%	78	Microscopy	Ecuador	(Martin-Solano et al., 2017) [121]	Primate
Strongyloides spp.	17%	53	Microscopy	Uganda	(Matsubayashi et al., 1992) [122]	Primate
Strongyloides fuelleborni	58%	432	Molecular	Uganda	(McLennan et al., 2017) [123]	Primate
Strongyloides spp.	84%	121	Microscopy	Uganda	(Muehlenbein et al., 2005) [124]	Primate
Strongyloides fuelleborni	45%	315	Microscopy	Kenya	(Munene et al., 1998) [125]	Primate
Strongyloides fuelleborni	21%	297	Microscopy	Kenya	(Muriuki et al., 1998) [126]	Primate
Strongyloides spp.	76%	83	Microscopy	Costa Rica	(Parr et al., 2013) [127]	Primate
Strongyloides spp.	13%	366	Microscopy	Tanzania	(Petrasova et al., 2010) [128]	Primate
Strongyloides spp.	44%	130	Microscopy	Tanzania	(Petrzelkova et al., 2010) [129]	Primate
Strongyloides stercoralis	15%	86	Microscopy	Peru	(Phillips et al., 2004) [130]	Primate
Strongyloides spp.	43%	47	Microscopy	Gabon	(Pouillevet et al., 2017) [131]	Primate
Strongyloides fuelleborni	6%	125	Microscopy	Cameroon	(Pourrut et al., 2011) [132]	Primate
Strongyloides spp.	53%	55	Microscopy	Ghana	(Ryan et al., 2012) [133]	Primate
Strongyloides spp.	8%	420	Molecular	Mexico	(Solorzano-Garcia and de Leon 2017) [134]	Primate
Strongyloides fuelleborni	39%	243	Molecular	Thailand and Laos	(Thanchomnang et al., 2018) [135]	Primate
Strongyloides spp.	35%	283	Microscopy	India	(Tiwari et al., 2017) [136]	Primate
Strongyloides stercoralis	31%	135	Microscopy	Thailand	(Wenz-Mucke et al., 2013) [137]	Primate
Strongyloides spp.	24%	272	Microscopy	South Africa	(Wren et al., 2015) [138]	Primate
Strongyloides spp.	24%	332	Microscopy	South Africa	(Wren et al., 2016) [139]	Primate
Strongyloides fuelleborni and Strongyloides spp.	UNK	14	Molecular	Malaysian Borneo	(Frias et al., 2018) [140]	Primate
Strongyloides spp.	29%	64	Microscopy	Brazil	(De Souza et al., 2012) [66]	Sheep
Strongyloides spp.	8%	165	Microscopy	Papua New Guinea	(Koinari et al., 2013) [141]	Sheep
Strongyloides spp.	<1%	27	Microscopy	New England	(MacGlaflin et al., 2011) [142]	Sheep
Strongyloides spp.	UNK	1798	Microscopy	Brazil	(McManus et al., 2009) [143]	Sheep
Strongyloides spp.	2%	275	Microscopy	Greenland	(Andreassen et al., 2017) [144]	Fox
Strongyloides spp.	4%	22	Microscopy	Iran	(Dalimi et al., 2006) [145]	Fox
Strongyloides stercoralis	16%	249	Microscopy	Mexico	(Hernandez-Camacho et al., 2011) [146]	Fox
Strongyloides stercoralis	2%	1198	Microscopy	Slovakia	(Miterpakova et al., 2009) [147]	Fox
Strongyloides spp.	65%	60	Microscopy	Pakistan	(Afshan et al., 2013) [16]	Rat
Strongyloides spp.	97%	299	Microscopy	Brazil	(Carvalho-Pereira et al., 2018) [17]	Rat
Strongyloides spp.	40%	25	Microscopy	Brazil	(Lima et al., 2017) [97]	Rat
Strongyloides spp.	13%	76	Microscopy	Bangladesh	(Fuehrer et al., 2012) [18]	Rat
Strongyloides spp.	10%	502	Microscopy	Nigeria	(Isaac et al., 2018) [19]	Mouse and rat
Strongyloides stercoralis	53%	98	Microscopy	Indonesia	(Prasetyo et al., 2016) [20]	House rat
Strongyloides spp.	10%	10	Microscopy	Brazil	(Souza et al., 2015) [21]	Capybaras
Strongyloides spp.	10%	31	Microscopy	Brazil	(Gioia-Di Chiacchio et al., 2014) [22]	Capybaras
Strongyloides stercoralis	2%	6530	Microscopy	Ethiopia	(Fetene and Worku 2009) [23]	Flies
Strongyloides stercoralis	<1%	9950	Microscopy	Ethiopia	(Getachew et al., 2007) [24]	Flies
Strongyloides stercoralis	2%	5000	Microscopy	Nigeria	(Umeche 1989b) [25]	Flies
Strongyloides stercoralis	12%	749	Microscopy	Nigeria	(Adenusi et al., 2018) [26]	Cockroaches
Strongyloides stercoralis	1%	920	Microscopy	Thailand	(Chamavit et al., 2010) [27]	Cockroaches
Strongyloides stercoralis	81%	70	Microscopy	Nigeria	(Morenikeji et al., 2016) [28]	Cockroaches
Strongyloides stercoralis	UNK	234	Microscopy	Nigeria	(Tatfeng et al., 2005) [29]	Cockroaches
Strongyloides stercoralis	2%	125	Microscopy	Nigeria	(Adesewa and Morenikeji, 2017) [82]	Soil
Strongyloides spp.	3%	625	Microscopy	Spain	(Dado et al., 2012) [38]	Soil
Strongyloides spp.	8%	120	Microscopy	Egypt	(Etewa et al., 2016) [83]	Soil
Strongyloides stercoralis	1%	797	Microscopy	Nigeria	(Ivoke et al., 2017) [85]	Geophagy
Strongyloides stercoralis	2%	1078	Microscopy	Tanzania	(Kawai et al., 2009) [86]	Geophagy
Strongyloides stercoralis	3%	112	Microscopy	Iran	(Motazedian et al., 2006) [39]	Soil
Strongyloides spp.	7%	182	Microscopy	Malaysia	(Noor Azian et al., 2008) [40]	Soil
Strongyloides stercoralis	20%	102	Microscopy	Nigeria	(Ogbolu et al., 2011) [41]	Soil
Strongyloides spp.	5%	2520	Microscopy	Brazil	(Rocha et al., 2011) [42]	Soil
Strongyloides spp.	2%	500	Microscopy	Czech Republic	(Valkounova 1982) [43]	Soil
Strongyloides spp.	3%	125	Microscopy	Brazil	(Mandarino-Pereira et al., 2010) [44]	Soil
Strongyloides stercoralis	14%	14	Microscopy	Slovakia	(Strkolcova et al., 2017) [45]	Soil
Strongyloides stercoralis	12%	17	Microscopy	South Africa	(Sumbele et al., 2014) [84]	Soil
Strongyloides spp.	4%	45	Microscopy	Romania	(Tudor 2015) [46]	Soil
Strongyloides stercoralis	6%	150	Microscopy	Nigeria	(Umeche 1989a) [47]	Soil
Strongyloides spp.	6%	16	Microscopy	Brazil	(da Silva et al., 2014) [148]	Soil
Strongyloides spp.	UNK	8	Microscopy	Cameroon	(Aghaindum and Landry, 2019) [149]	Non-potable water
Strongyloides spp.	40% - 100%	100	Microscopy	Saudi Arabia	(Bolbol 1992) [69]	Non-potable water
Strongyloides stercoralis	2%%	UNK	Microscopy	Brazil	(Bastos et al., 2008) [70]	Non-potable water
Strongyloides spp.	100%	3	Microscopy	Brazil	(Cutolo et al., 2006) [71]	Non-potable water
Strongyloides stercoralis	19%	52	Microscopy	Palestine	(Hilles et al., 2014) [150]	Seawater
Strongyloides stercoralis	1%	85	Microscopy	Turkey	(Bakir et al., 2003) [151]	Drinking water
Strongyloides fuelleborni and Strongyloides spp.	11% S. fuelleborni, 15% S. spp.	9950	Microscopy	Zimbabwe	(Dalu et al., 2011) [76]	Drinking water
Strongyloides spp.	UNK	UNK	Microscopy	Egypt	(El Shazly et al., 2003) [75]	Drinking water
Strongyloides stercoralis	7%	80	Microscopy	Egypt	(El-Badry et al., 2018) [152]	Drinking water
Strongyloides stercoralis	81%	16	Microscopy	Brazil	(Freitas et al., 2017) [153]	Drinking water
Strongyloides stercoralis	51%	232	Microscopy	India	(Jonnalagadda and Bhat 1995) [77]	Drinking water
Strongyloides stercoralis	100%	UNK	Microscopy	USA	(Klotz et al., 1992) [154]	Drinking water
Strongyloides stercoralis	UNK	UNK	Molecular	Malaysia	(Zeehaida et al., 2011) [155]	Fruit & vegetables
Strongyloides stercoralis	<1%	1130	Microscopy	Nigeria	(Adamu et al., 2012) [30]	Fruit & vegetables
Strongyloides stercoralis	<1%	960	Microscopy	Nigeria	(Adenusi et al., 2015) [31]	Fruit & vegetables
Strongyloides stercoralis	10%	150	Microscopy	Nigeria	(Amaechi et al., 2016) [78]	Fruit & vegetables
Strongyloides stercoralis	7%	190	Microscopy	Nigeria	(Amuta et al., 2017) [32]	Fruit & vegetables
Strongyloides stercoralis	7%	240	Microscopy	Nigeria	(Dada et al., 2015) [33]	Fruit & vegetables
Strongyloides spp.	1%	453	Microscopy	Iran	(Fallah et al., 2016) [34]	Fruit & vegetables
Strongyloides stercoralis	36%	360	Microscopy	Ghana	(Kudah et al., 2018) [35]	Fruit & vegetables
Strongyloides spp.	13%	108	Microscopy	Brazil	(Luz et al., 2017) [156]	Fruit & vegetables
Strongyloides spp.	19%	199	Microscopy	Nigeria	(Maikai et al., 2012) [157]	Fruit & vegetables
Strongyloides spp.	11%	36	Microscopy	Malaysia	(Matyusof et al., 2017) [80]	Fruit & vegetables
Strongyloides stercoralis	1%	260	Microscopy	Sudan	(Mohamed et al., 2016) [36]	Fruit & vegetables
Strongyloides stercoralis	10%	265	Microscopy	Thailand	(Punsawad et al., 2019) [37]	Fruit & vegetables
Strongyloides stercoralis	46%	120	Microscopy	Nigeria	(Ogbolu et al., 2009) [81]	Fruit & vegetables
Strongyloides stercoralis	14%	140	Microscopy	Iran	(Madadi 2010) [158]	Fruit & vegetables
Strongyloides stercoralis	19%	80	Microscopy	Nigeria	(Ohaeri and Unogu 2011) [79]	Fruit & vegetables
Strongyloides spp.	7%	15	Microscopy	Zambia	(Berentsen et al., 2012) [159]	Other animals
Strongyloides spp.	5%	272	Microscopy	Nepal	(Bista et al., 2017) [160]	Other animals
Strongyloides spp.	100%	1	Microscopy	Brazil	(Cardia et al., 2016) [161]	Other animals
Strongyloides spp.	4%	432	Microscopy	Spain	(Cordon et al., 2008) [162]	Other animals
Strongyloides spp.	40%	52	Microscopy	Russia	(González et al., 2007) [163]	Other animals
Strongyloides spp.	2%	956	Microscopy	India	(Gupta et al., 2018) [164]	Other animals
Strongyloides spp.	<1%	1005	Microscopy	Germany	(Hallinger et al., 2018) [165]	Other animals
Strongyloides spp.	31%	42	Microscopy	Japan	(Hasegawa et al., 2017) [166]	Other animals
Strongyloides spp.	<1%	400	Microscopy	Croatia	(Hermosilla et al., 2017) [167]	Other animals
Strongyloides spp.	64% - 99%	990	Microscopy	Mexico	(Hu et al., 2018) [168]	Other animals
Strongyloides spp.	4%	821	Microscopy	China	(Huang et al., 2014) [169]	Other animals
Strongyloides spp.	15%	2280	Microscopy	Pakistan	(Khan et al., 2010) [67]	Other animals
Strongyloides spp.	UNK	6	Microscopy	Namibia	(Kumba et al., 2003) [170]	Other animals
Strongyloides spp.	36%	58	Microscopy	Poland	(Mizgajska-Wiktor et al., 2010) [171]	Other animals
Strongyloides spp.	67%	12	Microscopy	Mexico	(Mukul-Yerves et al., 2014) [172]	Other animals
Strongyloides spp.	57%	201	Microscopy	Estonia	(Oja et al., 2017) [173]	Other animals
Strongyloides spp.	47%	383	Microscopy	Mexico	(Ojeda-Robertos et al., 2017) [174]	Other animals
Strongyloides spp.	7%	6	Molecular	Iberian Peninsula	(Perera et al., 2013) [175]	Other animals
Strongyloides spp.	3%	468	Microscopy	Poland	(Pilarczyk et al., 2015) [176]	Other animals
Strongyloides spp.	17%	86	Microscopy	Bangladesh	(Rahman et al., 2018) [177]	Other animals
Strongyloides spp.	3%	1883	Microscopy	Italy	(Rinaldi et al., 2009) [178]	Other animals
Strongyloides spp.	44%	163	Microscopy	Portugal	(Rosalino et al., 2006) [179]	Other animals
Strongyloides spp.	45%	82	Microscopy	Australia	(Turni and Smales 2001) [180]	Other animals
Strongyloides spp.	UNK	UNK	Microscopy	Namibia	(Turner et al., 2010) [181]	Other animals
Strongyloides spp.	UNK	UNK	Microscopy	Namibia	(Turner et al., 2012) [182]	Other animals
Strongyloides spp.	<1%	213	Microscopy	Kenya	(VanderWaal et al., 2014) [183]	Other animals
Strongyloides spp.	74%	243	Microscopy	Philippines	(Ybanez et al., 2018) [184]	Other animals

The most commonly identified reports of Strongyloides spp. were within primates (26% of all published works), and dogs (14% of all published works), as seen in Table A1. Other animals identified as environmental sources included cats, ruminants, rodents, and insects. Water, soil, as well as fruit and vegetables were all also identified as containing Strongyloides spp.

Fifty percent of all studies identifying Strongyloides spp. within primate populations identified the larvae to genus level only. S. fuelleborni was the next most frequently identified species at 40%. Parasitic infections were identified more frequently in terrestrial primates than arboreal primates [11,12]. Most studies (80%) employed microscopy, as seen in Table A1. Proximity to human populations and increased interaction with human populations was also frequently reported in infected populations [13,14]. Captive primates treated with anthelmintic drugs were also reported as carriers of Strongyloides spp. [15]. Sample size ranged from 7 to 3349, and prevalence within primate studies ranged from <1% to 100%, as seen in Table A1.

Domestic and stray dogs were the second most commonly identified source. Fourteen percent of all studies identifying Strongyloides spp. were within dogs, with sample sizes ranging from 35 to 879 and prevalence ranged from <1% to 45%, as seen in Table A1.

All studies reporting incidences of Strongyloides spp. within ruminant farming animals only identified Strongyloides to the genus level, as seen in Table A1.

Studies identifying rodents as a source of Strongyloides spp. accounted for 5% of the published works. Studies identified Rattus rattus, Rattus norvegicus, Mus musculus, Dasyprocta, and Hydrochoerus hydrochaeris as carriers of Strongyloides spp. [16,17,18,19,20,21,22]. Sample sizes for rodent-based studies ranged from 10 to 502. The prevalence ranged from 10% to 97%, as seen in Table A1.

Studies identifying insects within the order Diptera as a source of Strongyloides spp. accounted for 2% of the published works, as seen in Table A1. Identified insects within this order included flies of the genus Musca spp. and Lucilia spp. All studies identifying Strongyloides spp. within Diptera identified it from sites within the continent of Africa [23,24,25]. The sample sizes ranged from 5000 to 9950, and prevalence was between <1% and 2%.

Insects within the order Blattodea were identified in 2% of all studies, as seen in Table A1. Identified insects within this order include cockroaches from the genus Periplaneta spp. and Blattella spp. Four of the five identified studies reported Strongyloides spp. within populations of Blattodea in Africa. The remaining study identified Strongyloides spp. in Blattodea in Thailand. All studies identified infected insects within housing and food preparation areas [26,27,28,29]. The sample sizes of studies identifying insects within the order Blattodea ranged from 70 to 920, with the prevalence ranging from 1% to 81%.

Half (50%) of published works identifying parasitic contamination of vegetables and fruits found S. stercoralis upon leafy, rough-surfaced vegetables such as lettuces, cabbage, celery, spinach, and carrot [30,31,32,33,34,35,36,37]. The sample sizes for fruit and vegetable-based studies ranged from 36 to 1130, with prevalence ranging from <1% to 46%.

Countries where Strongyloides spp. was identified in soils in public areas included Spain, Iran, Malaysia, Nigeria, Brazil, the Czech Republic, Slovakia, and Romania, as seen in Figure 2 [38,39,40,41,42,43,44,45,46,47]. Geophagy, the purposeful consumption of soils, was also commonly identified as a factor in infection from soil-based sources.

Studies identifying environmental sources of S. stercoralis, S. fuelleborni, and Strongyloides spp. are distributed across the world. Areas with a large amount of research included Europe, Africa, and South East Asia, as seen in Figure 1. Areas lacking research include Oceania, and the Americas, as seen in Figure 1. Many published studies identified Strongyloides spp. within temperate regions as opposed to tropical regions, as seen in Figure 1.

Figure 1

Map displaying the global distribution of all reported environmental cases of Strongyloides stercoralis, Strongyloides fuelleborni, and Strongyloides spp. Where circles are representative of Strongyloides stercoralis, diamonds are representative of Strongyloides fuelleborni, and stars are representative of Strongyloides spp. The size of each shape is mapped to the number of studies published in that country. Location of shapes does not represent exact location of study, but country in which the study was completed. Colored fill of shapes was assigned to a single source and is consistent across all helminth species.

Microscopy was the most commonly used identification technique (90%). However, molecular detection was more common in recent publications. For example, in 2018, 6 of the 14 papers identified employed molecular-based techniques; however, in 2011, 1 of 11 papers published used molecular-based techniques, as seen in Table A1.

3. Discussion

3.1. Animals

Primates and domesticated or feral dogs (canids) adapt well to association with human settlements and cohabitation with humans, indicating the potential for transmission to humans. Contamination with feces from domesticated or synanthropic primates and dogs may lead to other environmental sources, such as water and soil, becoming reservoirs of Strongyloides spp. capable of causing infection. Most studies found in this review were based on primate and dog investigation, suggesting that these animals preferentially live closely with and benefit from humans. This habitual closeness presents a chance for environmental transmission of Strongyloides spp.

3.1.1. Canids

Studies that report parasites found in canid feces frequently investigate multiple parasites such as Ancylostoma spp., Giardia spp., and Strongyloides spp. These studies have often found low levels of S. stercoralis within otherwise highly parasitically infected populations [40,48,49,50,51,52,53,54,55,56]. Infection occurs more frequently in canids when they are living stray. This might be a result of exposure to infective Strongyloides spp. larvae occurring more frequently to these dogs than dogs living within homes [57]. Mass drug administration (MDA) to stray dogs has been implemented successfully for the control of rabies; accordingly, it may be an option for the control of Strongyloides spp. [10]. Isolated or infrequent anthelmintic treatment increases infection rates and so considered treatment must be implemented [10,58]. Studies identifying canid feces as containing Strongyloides spp. commonly also screened the samples for other parasites. Sample sizes ranged from 35 to 3465. The highest prevalence was reported by Beknazarova et al. [59] who screened 35 canine fecal samples from Australia, of which 49% were positive for Strongyloides spp. This low sample size with a high positivity rate in comparison with other studies is representative of the inconsistent fecal shedding of Strongyloides spp. as well as the endemic location of the study. The lowest prevalence was reported by Ardelean et al. [60] with 1% of 3465 samples positive from dogs within Romania. Strongyloidiasis was observed most commonly in dogs three to six months of age in this study. This variance based on age and study location may be further impacted by the detection method. Ardelean et al. [60] reported high levels of Ancylostomidae spp. which is morphologically similar to Strongyloides spp., therefore making reliable identification with microscopy alone difficult.

3.1.2. Primates

Areas sparsely populated by humans increase roaming in primates due to the attractive food sources but offer a low threat from the decreased human numbers. More frequent entry to communities in search of food potentially increases the numbers of Strongyloides spp.-infected primate feces within these sparsely populated communities [13,14]. Terrestrial Papio primates were likely to excrete Strongyloides spp. larvae; however, arboreal Cercopithecus neglectus were less likely [11,12]. This may be due to less frequent contact with soil containing Strongyloides spp. larvae. The impact of human populations upon forests has led to an increased chance of interaction between humans and potentially infected primates. Hasegawa et al. [61] observed that degraded forest increased the chance of roaming and transfer of parasites.

Captive primates present an infection risk to handlers because anthelmintic treatment has been observed to not eliminate Strongyloides spp. larvae shedding within feces [15]. This may be due to the introduction of new individuals to groups, a phenomenon also observed within wild individuals [15]. This indicates the value in introducing physical environmental controls beyond anthelmintic drugs, especially in communities exposed to roaming wild primates.

Tourist sanctuaries provide an ideal environment for contact between primates and humans. Environmental controls such as fecal contamination removal can decrease helminthic infection in both primates and humans without interfering with natural behaviors [62]. Strongyloides spp. is unable to transfer either from animal to human or from human to human directly [63]. This further supports the importance of clearing feces because contact with the animals does not cause infection; however, contact with fecal matter can cause infection. Larger groups, such as those within tourist sanctuaries, are generally associated with higher parasitic species richness. Some variation of infection can be expected based on food availability and stress levels [64].

Studies of primates had sample sizes ranging from 7 to 3349, with prevalence also ranging from <1% to 100%. Prevalence within primate populations was reported to be higher than in canine populations. Hasegawa et al. [61] reported 100% prevalence within seven gorilla and chimpanzees from Uganda; whereas Li et al. [65] screened 3349 fecal samples and identified a prevalence rate of 6%. This variation in prevalence may be due to the inconsistent shedding of Strongyloides spp. larvae. Li et al. [65] employed microscopy whereas Hasegawa et al. [61] employed molecular techniques, which may account for differentiation in prevalence.

3.1.3. Ruminants

Strongyloides spp. has also been found within the feces of ruminants used in western farming settings including pigs, sheep, and cattle. All studies identifying Strongyloides spp. within farm-associated ruminants only identified the parasite to the species level [66,67]. These may have been genus specific, such as the pork threadworm, Strongyloides ransomi, or the more general Strongyloides papillosis. All studies used microscopy, a technique that can have low success in identifying Strongyloides spp. to species level. These recorded observations indicate the potential for infected ruminant feces to provide an environmental source of Strongyloides spp.

3.1.4. Rodents

Rodents are known to carry a range of communicable diseases. Strongyloides spp. has been found in several rodent species, including common house rats, and non-synanthropic rodents such as Hydrochoerus hydrochaeris. S. stercoralis has been identified in house rat feces in East Java, Indonesia, using microscopy [20]. The area in which S. stercoralis was identified in house rat feces is an area with poor sanitation and hygiene. People reported a large house rat population within these areas [20]. In such cases, where a zoonotic pathogen is identified, control of the offending carrier can be employed. The sample sizes of rodents were low in comparison with other sources, with the highest sample size being 502 [19]. The highest prevalence was within a population of Rattus norvegicus within a Brazilian slum [17]. The rodents sampled within this study had particularly high levels of infection with helminths; in all except five, helminths were present within their feces [17]. This prevalence of 97% from a sample size of 299 is representative of animals living within an area highly contaminated by human waste.

3.1.5. Insects

Increasing urbanization has allowed for synanthropic dependence to increase within insect populations. Densely urbanizing areas lead to an increase in available food for insects, and areas with poor sanitation and hygiene practices attract disease-carrying insects such as those within the order Blattodea (cockroaches) and Diptera (flies). Filth flies present a source of helminth transmission. Their preference for consuming wet, rotting substances indicates a high probability for the consumption and carriage of Strongyloides spp. Carriage of Strongyloides spp. has been observed on the external body of flies despite frequent preening and cleaning [68]. Fetene and Worku [23] identified S. stercoralis within Chrysomya rufifacies, Musca sorbens, and Lucilia cuprina. C. rufifacies were identified largely within butcheries and defecating grounds; M. sorbens was found more frequently within the market collection sites. Furthermore, Musca domestica, a species always found in association with humans, has been observed to carry Strongyloides spp. [24]. The presence of these flies within human food areas presents a potential transmission route for Strongyloides spp. larvae. Prevalence was higher in the internal structures of flies than on the external surface of flies [25]. This observation is further supported by the preference of these flies for consumption of wet substances. Insects within the order Blattodea, commonly known as cockroaches, also present a transmission source. Parasite prevalence has been found to be associated with housing type. Low-cost housing with pit latrines as well as housing in close proximity to dumpsites was reported to contain higher levels of carrier cockroaches [26,28,29]. Through the introduction of environmental controls such as fly screens or nets, movement of carrier insects can be decreased [27]. Sample sizes for both orders were high. Studies found low prevalence with the exception of Morenikeji et al. [28] who reported 81% in 70 cockroaches.

3.2. Water

Contamination of water is also a potential source of helminth transferal. Pollution of water sources with human and agricultural waste can render water sources unsuitable for use as drinking and irrigation water. In areas where water access is limited, contaminated water may be employed for these uses [69]. Waste stabilization ponds, chlorination, or activated sludge treatment systems may be suitable approaches for reducing helminth levels; however, many studies monitoring wastewater treatment methods have provided contradictory results [70,71,72,73]. Some studies identified standard treatment techniques as adequate for removing larvae; however, others did not. Frequent monitoring of treated waste water is important because treated water has been identified as containing higher than acceptable levels of helminths including Strongyloides spp. [74]. To date, studies have focused on the helminth burden of treated water instead of comparing treated with untreated levels. Through focusing on untreated and treated waters from the same area, reduction in burden levels of treated water may be better understood.

Untreated water used for drinking can contain Strongyloides spp., particularly when water is sourced from storm water or collected rainwater [75]. According to one study, when water runoff moves into drinking water sources such as rivers, it can carry Strongyloides spp. larvae with it [75]. Bore and ground-water contamination can also occur and has been identified [76,77]. Jonnalagadda and Bhat [77] found that improper washing of vessels used to collect and store water can lead to helminth contamination. Implementation of appropriate washing and sanitation education in areas with high contamination risk may decrease incidences of infection.

Prevalence of Strongyloides spp. within non-potable water was higher than in potable water; this was expected because most sources of non-potable water were collected from wastewater treatment facilities [71]. The prevalence of Strongyloides spp. within non-treated wastewater was between 40–100%. Treated waste water intended for use on crops for human consumption had a much lower prevalence (2%); however, it was still observed to be present, supporting the importance of monitoring water intended for reuse [70].

3.3. Fruit and Vegetables

The rough nature of green, leafy vegetables surfaces means that adhesion of parasitic larvae and eggs occurs easily when these vegetables are either washed with contaminated water or come into contact with contaminated human fecal-based fertilizers (i.e., night soil) [30]. Studies identified S. stercoralis contamination most frequently within leafy, rough-surfaced vegetables such as lettuces, cabbage, celery, spinach, and carrot [30,31,32,33,34,35,36,37]. This correlation may be due to these vegetables growing close to or in the ground, which may lead to increased contamination from fertilizers [78,79]. Market vendors commonly wash vegetables prior to purchase, and consumption of raw vegetables such as salad leaves is frequently noted [31,79,80]. There is an increasing focus on the study of vegetables, washing water, and farm soil to determine where in the food chain parasites are being introduced [33].

Prevalence of Strongyloides spp. within fruit and vegetable samples was generally low, ranging from <1% to 46%. Ogbolu et al. [81] found S. stercoralis in 46% of fresh vegetables sold at open markets in Nigeria. The application of night soils and untreated wastewater is common within low-income nations may have led to the high level of prevalence [81]. Lower prevalence was also reported within Nigeria, between <1% and 19%. This variation may be due to differences in handling of samples, treatment during farming, and cross contamination [30,31].

3.4. Soil

Increasing urbanization has led to an ever-increasing amount of waste. Modern waste includes not only fecal waste but waste produced in the form of rubbish. Dumpsites and landfills are commonly employed to deal with this large amount of waste; Strongyloides spp. contamination can occur throughout the nearby environments. Dumps and landfills pose a transmission risk due to the ability of Strongyloides spp. larvae to survive effectively in the soil [82].

Contamination of soils with animal feces within public recreation areas also presents a transmission source. High levels of soil-transmitted helminths were reported in public area soils such as parks in Spain, Iran, Malaysia, Nigeria, Brazil, the Czech Republic, Slovakia, and Romania [38,39,40,41,42,43,44,45,46,47].

The texture and chemistry of soil also plays a role in the prevalence of Strongyloides spp. larvae. Moisture levels in soils increases the incidence of rhabditiform larvae developing into filariform larvae [83]. This requirement for moisture is supported by the findings of a study of helminth larvae during the wet season, which found that no larvae were located during the dry season despite contamination [39]. High sand and silt content soils favor the survival of Strongyloides spp. and other helminth larvae. This is due to the high porosity of these soils, which allows larvae to move effectively through the soils towards sources of nutrition and moisture [84].

Strongyloides spp. is transmitted from soil-based sources to humans through skin-to-soil contact; however, the behavior of purposefully ingesting soil known as geophagy can also lead to soil-based infections. Geophagy is culturally accepted and common in sub-Saharan Africa. This behavior is common in pregnant women; S. stercoralis infections have been observed, along with other soil-transmitted helminth infections, in these women [85,86]. Geophagy may be undertaken as a method for diet supplementation in low-income areas. Notably, these areas are more likely to have helminth-contaminated soil, which leads to an increased chance of infection.

Prevalence of Strongyloides spp. within soil was between 1% and 20%. Ivoke et al. [85] screened 797 pregnant women for parasitic infections related to geophagy. The prevalence of infection was 1% within this cohort. This low prevalence is likely due to picking soils specifically for consumption. Higher prevalence was observed in soils sampled in soil directly from areas densely populated with poor health infrastructure [41].

3.5. PCR and Microscopy

Identification of Strongyloides spp. nematodes can be undertaken using several methods. These fall into either techniques involving the identification of Strongyloides spp. larvae using a microscope or molecular-based techniques. Microscopy presents several problems but it accounts for most larval identification-based techniques (90%) as presented in this literature review. This study did not exclude papers based on year; accordingly, this overrepresentation of microscopy is likely a result of the recent introduction of polymerase chain reaction (PCR) techniques. Recent papers have increasingly employed PCR as accessibility to the equipment increases. In 2018, 5 of the 14 papers published employed molecular-based techniques; in contrast, in 2011, only 1 of 11 studies published employed molecular-based techniques, as seen in Table A1. This increasing use of molecular-based techniques is expected, and its use will allow for more accurate identification of species of Strongyloides spp. The strengths of microscopy include that it can be employed within the field or where resources are limited; however, microscopy alone cannot reliably differentiate S. stercoralis from S. fuelleborni. The reliance on microscopy-based techniques is hazardous because both species are morphologically similar [34]. Molecular techniques allow for the accurate identification of Strongyloides spp. to the species level; however, set up of molecular protocols can be expensive. Each technique has strengths and weaknesses; when looking at all published works, a consideration of the identification techniques allows for more accurate assessment of reports.

3.6. Global Distribution

Globally, published research has mainly focused in countries within Africa, Europe, and South East Asia. Commonly, Strongyloides spp. is reported as a tropical disease; however, Strongyloides spp. was often also reported in temperate regions such as Europe, as seen in Figure 1 [83,87]. This may be due to a lack of resources within low-income nations, leading to an overrepresentation of the generally higher income countries within Europe. Australia and the Americas both lacked studies looking at the environmental sources of Strongyloides spp. (Figure 1). This indicates a need for more research into the environmental transmission of S. stercoralis, S. fuelleborni, and Strongyloides spp. within these areas.

4. Materials and Methods

This systematic literature review is based on an adapted version of the PRISMA statement. This tool allows for the transparent and reliable reporting of evidence. A systematic search of the databases Scopus and Web of Science was undertaken, and all articles published prior to 2019 were included. Key words used in searches included Strongyloides spp., strongyloidiasis, tap water, soil, insect, zoonotic, and waste, as seen in Table 1. A search strategy was developed to ensure a transparent and complete literature review of all identified environmental sources of Strongyloides spp. was completed. This strategy is as follows; All non-English documents were excluded from the search.

To be included, published data must have reported Strongyloides spp. in one of the three spp. capable of human infection or to the genus level because these studies cannot be excluded as identifying disease-causing Strongyloides spp. The document must have reported this presence within an environmental source. Documents were excluded if they were reviews, reports of humans with Strongyloides spp. infection with no mention of contributing environmental source, or lab-based studies, as seen in Figure 2.

Figure 2

Flow diagram representing the search strategies used (based on the PRISMA statement reporting guidelines for systematic literature reviews) showing an overview of the retrieved articles and the total articles identified as eligible.

First, all titles and abstracts of all papers were manually reviewed to ensure the papers met inclusion criteria. If it was unclear from titles and abstracts if papers met the criteria, they were included for full text review. Papers that were unclear were included. Papers were then read as full text and compared against inclusion and exclusion criteria. Articles that met these criteria were included in the study. All papers included in the study had key points extracted and recorded including the environmental source reported, species of Strongyloides spp. observed, detection method used, and the country from which the sample was taken.

5. Conclusions

Although Strongyloides spp. is considered a soil-transmitted helminth, there are several environmental sources that can potentially provide a route of transmission of the disease. Understanding the potential sources, combined with the adoption of environmental controls for Strongyloides spp. is likely to decrease transmission and therefore infections. Animals such as dogs, primates, and insects, as well as soil, water, and fruit and vegetables have all been reported to contain Strongyloides spp. larvae, capable of perpetuating infection within humans who have come into contact. Future research is needed to undertake a holistic screening of all environmental sources within endemic areas to identify those which pose the greatest significance to human health. By understanding the established and recorded environmental reservoirs of S. stercoralis, S. fuelleborni, and S. fuelleborni kellyi, better environmental controls can be implemented.