Transgenesis and targeted mutagenesis in the human-parasitic nematode Strongyloides stercoralis

Introduction

Soil-transmitted helminths (STHs) are a classification of parasitic nematodes characterized by transmission via ingestion of infective eggs or larvae, or by contact with infective larvae in soil contaminated with feces (Buonfrate et al., 2023; Czeresnia and Weiss, 2022; Loukas et al., 2021). STHs include hookworms in the genera Necator and Ancylostoma, the giant intestinal roundworm Ascaris lumbricoides, the whipworm Trichuris trichiura, and the human threadworm Strongyloides stercoralis (Buonfrate et al., 2023; Czeresnia and Weiss, 2022; Loukas et al., 2021). These gastrointestinal helminths infect over 1.5 billion people worldwide, causing chronic debilitating conditions, and S. stercoralis infections can be fatal in immunocompromised individuals (Buonfrate et al., 2020; Chen et al., 2024; Fleitas et al., 2022; Loukas et al., 2021; Riaz et al., 2020). In addition, children infected with STHs may have stunted growth or nutrient deficiencies, resulting in life-long medical complications (Fauziah et al., 2022; Forrer et al., 2017). Economically disadvantaged individuals living in tropical or subtropical regions with inadequate access to proper sewage and health infrastructure are disproportionately impacted by STHs (Beknazarova et al., 2016; Loukas et al., 2021; Lynn et al., 2021; Magalhães et al., 2023; McKenna et al., 2017). Individuals that come in contact with STH-contaminated soil are infected either via the fecal-oral route (A. lumbricoides and T. trichiura) or skin penetration (hookworms and S. stercoralis) (Buonfrate et al., 2023; Loukas et al., 2021). Some hookworms, and possibly S. stercoralis, can also infect when infective larvae are ingested (Buonfrate et al., 2023; Loukas et al., 2021). While ivermectin is administered as a gold-standard anthelmintic treatment, there are currently no prophylactics or vaccines available against any of the STHs, and subpar drug efficacy and resistance are major concerns (Buonfrate et al., 2023; Coffeng et al., 2024; Fissiha and Kinde, 2021; Ng’etich et al., 2023; Sengthong et al., 2021).

S. stercoralis is the causative agent of the neglected tropical disease strongyloidiasis, which infects an estimated 300–600 million people globally (Buonfrate et al., 2020; Fleitas et al., 2022). Individuals infected with S. stercoralis may experience gastrointestinal distress. However, severe cases arise when individuals experience hyperinfection (high worm burden) or larval dissemination throughout the body, typically as a result of immunosuppression (Buonfrate et al., 2023). Efforts to control infections are complicated in cases of autoinfection, where the parasites chronically reinfect the same host. Treatment for autoinfection requires repeated administration of anthelminthics over week-long intervals; coupled with lack of reliable rapid detection methods, this makes strongyloidiasis a particularly challenging disease to eradicate (Buonfrate et al., 2023; Lo et al., 2025; Luvira et al., 2022; Tamarozzi et al., 2023). Efforts to eliminate skin-penetrating nematodes include improving local sanitation infrastructure, wearing closed-toed shoes, and extensive surveillance programs. However, these methods alone are not sufficient to reduce chronic or disseminated infections (Buonfrate et al., 2023; Lo et al., 2025). Thus, understanding the molecular mechanisms that drive parasite pathogenesis is critical for optimizing the development of targeted prophylactics, diagnostics, and therapeutics.

S. stercoralis has a complex life cycle that consists of both free-living and parasitic stages (Figure 1A). Developmentally arrested soil-dwelling infective third-stage larvae (iL3s) locate human hosts using host-emitted sensory cues, including body heat and odorants (Bryant et al., 2018, 2022; Castelletto et al., 2014; Ekobol et al., 2025; Gang et al., 2020; Lopez et al., 2000; Safer et al., 2007). The parasitic life cycle begins when the iL3s release a protease-rich excretory-secretory product and burrow through host skin (Czeresnia and Weiss, 2022; Dishnica et al., 2023; Gomez Gallego et al., 2005; Hunt et al., 2016; McKerrow et al., 1990; Moser and Hallem, 2024). The iL3s then resume development, a process called activation, and migrate through the host circulatory and respiratory systems until ultimately colonizing the crypts of the small intestine as parthenogenetic parasitic adult females (Buonfrate et al., 2023). The progeny of the parasitic adult females can either: 1) develop into larvae that reinfect the same host in a process called autoinfection; 2) exit the host in feces as L1 larvae and develop directly into iL3s; and/or 3) exit the host and undergo a single free-living generation consisting of sexually reproductive adult males and females, whose progeny all develop into iL3s (Buonfrate et al., 2023) (Figure 1A).

S. stercoralis free-living adults are amenable to transgenesis and targeted mutagenesis using techniques for intragonadal microinjection of exogenous DNA adapted from those used in the free-living model nematode Caenorhabditis elegans (Figure 1B) (Castelletto et al., 2020; Mendez et al., 2022). These methods are enabled by the sequencing and annotation of the S. stercoralis genome, as well as the generation of high-quality RNA-sequencing datasets (Howe et al., 2017; Hunt et al., 2018, 2016, 2017). Here, we review the development of techniques for transgenesis and targeted mutagenesis in S. stercoralis. We also discuss how recent technical advances have enabled the generation of the first S. stercoralis transgenic and knockout stable lines (Banerjee et al., 2025a; Patel et al., 2025, 2024), thereby greatly facilitating comprehensive studies of gene function in these parasites.

C. elegans as a comparative model for S. stercoralis

As a well-established genetic model system, the free-living nematode C. elegans offers a powerful starting point for understanding the biology of S. stercoralis and other parasitic nematodes. For example, in C. elegans, insulin signaling via the insulin receptor Dauer Formation-2 (DAF-2) regulates the entry into and exit from the dauer larval stage, a developmentally arrested third-larval stage that is homologous to the iL3 stage of parasitic nematodes (Hu, 2007; Viney et al., 2005). S. stercoralis has a daf-2 homolog, and aptazyme-mediated disruption of Sst-DAF-2 impairs resumption of development by iL3s upon exposure to host-like conditions (Zhang et al., 2025). Thus, insulin signaling regulates larval development in both C. elegans and S. stercoralis. Similarly, carbon dioxide (CO₂) detection in both C. elegans and S. stercoralis requires the receptor guanylate cyclase GCY-9, a putative receptor for CO₂, and the BAG sensory neurons (named for their bag-like dendritic endings) (Banerjee et al., 2025a; Hallem et al., 2011; Smith et al., 2013). However, it is important to note that C. elegans and S. stercoralis are distantly related species from different nematode clades, and there is often not a one-to-one correspondence between genes in C. elegans and S. stercoralis (Ahmed et al., 2022; Blaxter et al., 2016, 1998). For example, the astacin family of zinc metalloproteases is present in both C. elegans and S. stercoralis; however, while C. elegans has 40 astacins, S. stercoralis has over 200 (Hunt et al., 2016, 2017; Möhrlen et al., 2003; Moser and Hallem, 2024; Park et al., 2010). The expansion of the astacin family in S. stercoralis suggests an important role for these proteins in nematode parasitism (Hunt et al., 2016, 2017; Moser and Hallem, 2024) and highlights the limitations of relying entirely on C. elegans to inform studies of parasitic nematodes. The direct study of parasitic nematodes is critical for uncovering the specific molecular and cellular adaptations that enable them to successfully parasitize human hosts.

Fortunately, many of the molecular genetic tools used to engineer transgenic and mutant nematodes originally developed in C. elegans have proven to be adaptable to S. stercoralis. In C. elegans, transgenic and knockout lines can be generated by microinjection of exogenous DNA into the syncytial gonad of young adult hermaphrodites, where it is then taken up into developing oocytes (Evans, 2006; Kim et al., 2022). In the case of transgenesis, both linear and plasmid DNA can be incorporated into extrachromosomal arrays in the F₁ progeny, which are then inherited transgenerationally (Evans, 2006). In the case of CRISPR-mediated genome editing, either DNA plasmids encoding the single guide RNA and Cas9 endonuclease, or a ribonucleoprotein complex consisting of the Cas9 protein and single guide RNA, are introduced into adult hermaphrodites to generate heritable mutations in the F₁ progeny (Kim et al., 2022). In the absence of a repair template, Cas9-mediated double-stranded breaks are repaired by non-homologous end joining, typically resulting in small indels that disrupt gene function. However, if DNA containing a repair template is included in the microinjection mix, then the double-stranded break can be precisely repaired by homology-directed repair, resulting in incorporation of the DNA sequence contained in the repair template (Kim et al., 2022). Uses of CRISPR-mediated genome editing include disrupting or editing genes, endogenously tagging genes, and inserting transgenes through targeted integration into the genome (Dickinson and Goldstein, 2016; El Mouridi et al., 2022; Kim et al., 2022).

Transgenesis in S. stercoralis

Like C. elegans adults, S. stercoralis free-living adults have a syncytial gonad; thus, exogenous DNA can be introduced by intragonadal microinjection using techniques adapted from C. elegans (Figure 2A) (Castelletto and Hallem, 2021; Junio et al., 2008; Li et al., 2006; Shao et al., 2017). The generation of transgenics via intragonadal microinjection is not yet feasible in parasitic nematodes such as hookworms that lack a free-living (and therefore easily accessible) adult life stage. However, heritable transgenesis via intragonadal microinjection has been successfully applied to the rat-parasitic nematode Strongyloides ratti and the possum-parasitic nematode Parastrongyloides trichosuri, both of which have a life cycle that includes a free-living adult stage similar to that of S. stercoralis (Grant et al., 2006; Li et al., 2011; Lok, 2012; Lok et al., 2017; Shao et al., 2012). Successful transgenesis requires the use of endogenous promoters and regulatory elements (Junio et al., 2008; Li et al., 2006), and transgenesis rates are greatly improved by the codon optimization of transgenes (Bryant and Hallem, 2021; Patel et al., 2024). It is important to note that manual curation of S. stercoralis genes is also often critical to ensure appropriate transgene design (Bryant et al., 2024).

While C. elegans can express transgenes from extrachromosomal arrays across generations (Evans, 2006), this feature is not shared with S. stercoralis. In Strongyloides, extrachromosomal arrays are silenced after the F₁ generation (Junio et al., 2008; Lok, 2013; Shao et al., 2012). Thus, heritable expression in Strongyloides requires genomic integration of transgenes. This has been achieved using the piggyBac system (Lok, 2013, 2012; Shao et al., 2012, 2017). The piggyBac transposase, originally derived from the cabbage looper moth Trichoplusia ni (Yusa, 2015), drives chromosomal integration by excising DNA sequences flanked by inverted terminal repeats (ITRs) and integrating them into the genome at sites containing TTAA motifs (Sandoval-Villegas et al., 2021). To integrate transgenes using the piggyBac system, two plasmids are microinjected: one encodes the transgene of interest flanked by ITRs and the other encodes the piggyBac transposase. Advantages of the piggyBac system include the ability to integrate large transposons of over 100 kb and the high copy number of integrated transgenes, resulting in robust transgene expression (Sandoval-Villegas et al., 2021). In S. ratti, copy number estimates ranged from ~30–50 per genome (Shao et al., 2012).

Until recently, even with the use of piggyBac, the generation of stable transgenic lines that could be maintained indefinitely across generations was feasible with S. ratti but not S. stercoralis (Lok, 2012). This is because maintenance of S. stercoralis requires passaging infective larvae through a laboratory host such as the Mongolian gerbil (Meriones unguiculatus), and subcutaneous injection of 1,000-2,000 iL3s per gerbil was required to establish a patent infection (Nolan et al., 1993). The relatively low transgenesis rate of S. stercoralis (typically, transgenic larvae comprise <5% of the F₁ iL3 population) made generating ~1,000 transgenic F₁ iL3s impractical (Castelletto and Hallem, 2021). In one study, sufficient transgenic S. stercoralis F₁ iL3s were collected and used to successfully infect a gerbil; however, only two transgenic larvae were obtained in the F₂ generation, precluding further host passage (Lok, 2012). Thus, studies of transgenic S. stercoralis were limited to small numbers of transgenic F₁ iL3s.

These limitations were recently overcome to generate the first stable transgenic lines of S. stercoralis (Figure 2A) (Patel et al., 2024). Stable transgenesis was achieved through several modifications to the original transgenesis protocol: 1) A hyperactive piggyBac transposase, hyPBase, was used for genomic integration, since hyPBase was found to have a ~9-fold higher integration rate in mammalian cells (Yusa et al., 2011); 2) the hyPBase transgene was codon-optimized for expression in S. stercoralis; 3) transgenic iL3s were subjected to in vitro activation, a process whereby the iL3s are exposed to host-like conditions that cause them to initiate development outside the host (Gang et al., 2020; Stoltzfus et al., 2014, 2012); and 4) the iL3s were introduced into the gerbil by oral gavage instead of subcutaneous injection (Patel et al., 2024). Transgenic worms were then isolated from gerbil feces (Patel et al., 2024). Infecting gerbils with activated iL3s by oral gavage reduced the number of larvae required to establish an infection from ~1,000 to ~100, making it more easily feasible to obtain sufficient numbers of transgenic larvae by intragonadal microinjection (Patel et al., 2024). Using this approach, transgenic S. stercoralis, with stable transgenic phenotypes, can be propagated in gerbils across multiple generations. The generation of stable transgenic lines offers numerous experimental advantages, including the ability to propagate transgenic worms through gerbil hosts indefinitely, less mosaic expression patterns, and the ability to generate large numbers of transgenic worms of any life stage (Figure 2B) (Patel et al., 2024).

Transgenic S. stercoralis have been used extensively in many different studies. S. stercoralis iL3s expressing transcriptional reporter constructs, in which an endogenous promoter sequence is used to drive expression of a fluorophore gene, have been used to examine gene expression patterns in multiple contexts (Banerjee et al., 2025a; Bryant et al., 2018, 2022; Castelletto et al., 2009; Cheong et al., 2021; Lok, 2012; Patel et al., 2025, 2024; Stoltzfus et al., 2012; Zhou et al., 2023). Transgenic S. stercoralis have also been used to study gene function through expression of dominant negative alleles (Castelletto et al., 2009) and, more recently, through the use of aptazymes to generate conditional gene knockdowns (Zhang et al., 2025). Transgenesis has also been used to genetically target specific subsets of neurons for studies of neuronal function, including neuronal silencing coupled with quantitative behavioral analysis (Banerjee et al., 2025a; Bryant et al., 2022; Patel et al., 2025) and in vivo calcium imaging of neuronal activity (Banerjee et al., 2025a; Bryant et al., 2022). Finally, stable transgenic lines that express the fluorophore mScarlet-I in body-wall muscle (Figure 2B) have been essential for studies of skin penetration by enabling visualization of the normally translucent worms on the surface of the skin (Figure 2C) (Abell et al., 2025; Patel et al., 2025).

Although generating transgenic lines using the piggyBac system is straightforward and efficient, there are several disadvantages of this approach. First, integration with piggyBac occurs in a semi-random manner that has the potential to cause unintentional gene disruptions (Sandoval-Villegas et al., 2021; Yusa, 2015). Additionally, the number of transgene copies that are integrated into the genome cannot be controlled, making it impossible to mimic endogenous expression levels (Sandoval-Villegas et al., 2021; Yusa, 2015). Finally, because the transgenes integrate at high copy number, it is difficult to make the transgene homozygous at every integration site to obtain a true homozygous line. As a result, maintaining a stable transgenic line requires the periodic selection of bright transgenics to ensure that non-transgenic worms or low-expressing worms do not increase in population frequency over successive host passages. CRISPR/Cas9-mediated genomic integration, described below, offers a powerful alternative for targeted genomic integration in cases where lower copy number is required or less bright transgene expression is acceptable.

Targeted mutagenesis in S. stercoralis

S. stercoralis is also amenable to CRISPR/Cas9-mediated targeted mutagenesis using the same intragonadal microinjection technique that is used to make transgenics (Gang et al., 2017; Lok et al., 2017). Typically, plasmids encoding the single guide RNA(s) for targeting the gene of interest, the Cas9 endonuclease, and a template for homology-directed repair (HDR) that encodes a fluorescent reporter are microinjected into the syncytial gonad of free-living females; homozygous iL3s can then be found in the F₁ generation (Banerjee et al., 2025a; Bryant et al., 2018, 2022; Cheong et al., 2021; Gang et al., 2017, 2020). The Cas9 target sites that result in efficient editing are the same in S. stercoralis and C. elegans (Banerjee et al., 2025b; Farboud, 2017), making target site selection straightforward. Among the F₁ progeny from microinjected females, candidate mutants can be identified based on expression of the fluorescent marker, and from this population, homozygous mutants can be identified by single-worm PCR genotyping (Gang et al., 2017). CRISPR-induced mutations can also be obtained by injecting the CRISPR components as a ribonucleoprotein complex (RNP) (Adams et al., 2019; Gang et al., 2017); however, in at least some cases, homology-directed repair appears to be more efficient when plasmids are used instead of RNPs (Gang et al., 2017). Additionally, a template for homology-directed repair does not need to be included; however, in the absence of a repair template, large deletions rather than small indels have been observed (Gang et al., 2017).

In these earlier CRISPR studies, stable mutant lines were not generated. Rather, the F₁ iL3 progeny of microinjected females were screened for expression of the fluorescent reporter from the HDR construct, subjected to single-worm microscopy or behavioral analysis, and then genotyped post hoc to identify homozygous knockouts (Banerjee et al., 2025a; Bryant et al., 2018, 2022; Cheong et al., 2021; Gang et al., 2017, 2020). This approach was time-consuming and labor-intensive because a majority of the candidate mutants tested were not homozygous knockouts – some were heterozygous for the mutation, some were mosaic, and some were wild type but expressed the fluorescent reporter from an extrachromosomal array. Only a few homozygous knockouts could be obtained from a given microinjection experiment, and the CRISPR pipeline had to be repeated multiple times to obtain sufficient sample sizes.

More recently, an approach for developing the first stable knockout lines was developed (Figure 2D) (Banerjee et al., 2025a, 2025b; Patel et al., 2025), using a modification of the approach for making stable transgenic lines (Patel et al., 2024). To make stable knockout lines, two separate groups of free-living females are microinjected in parallel; one group receives a microinjection cocktail that includes an HDR construct with a red fluorophore and a second group receives a microinjection cocktail that includes an HDR construct with a green fluorophore (alternatively, a blue fluorophore can be used instead of the red or green fluorophore). The F₁ iL3 progeny from these two groups of microinjected females are screened for red or green fluorescence, and the red or green iL3s are then pooled. Following the same strategy used for generating stable transgenic lines (Patel et al., 2024), these red or green iL3s are then activated in vitro, pooled, and introduced into a gerbil host by oral gavage. Once the gerbil is patent, it will shed F₂ progeny, some of which will develop into red or green free-living adults. The free-living adults will then mate on feces to generate red/green dual-color F₃ iL3s. These dual-color iL3s are homozygotes, with a red fluorophore integrated into one allele of the target gene and a green fluorophore integrated into the other allele (Figure 2E). Notably, because extrachromosomal arrays are silenced after the F₁ generation in S. stercoralis, all fluorescence in subsequent generations reflects integrated transgenes, and thus all red/green worms are homozygous knockouts. These homozygotes can then be propagated by subsequent passages through a gerbil (Figure 2D). Alternatively, in cases where homozygotes show infection defects, the worms can be propagated through gerbil hosts as heterozygotes, and homozygotes can be obtained as needed by mating red free-living females with green free-living males, or vice versa (Banerjee et al., 2025b; Patel et al., 2025). With stable mutant lines, it is possible to obtain large numbers of homozygous mutant worms of any life stage. This approach was recently used to demonstrate a role for the receptor guanylate cyclase gene Sst-gcy-9 in mediating CO₂ response (Banerjee et al., 2025a), and to demonstrate a role for the tyrosine hydroxylase gene Sst-cat-2 in driving skin-penetration behavior (Figures 2E, F) (Patel et al., 2025). It should now be possible to use this approach to study nearly any non-lethal gene in S. stercoralis.

Conclusions and future directions

The ability to make stable transgenic and knockout lines of S. stercoralis has greatly accelerated mechanistic studies of these parasites. However, several important directions for future technical development remain. First, S. stercoralis cannot yet be maintained without an animal host, and host passage is both expensive and time-consuming. An in vitro culturing system would be invaluable for the low-cost maintenance of stable knockout and transgenic lines. Second, although most genes can now be targeted for mutagenesis using CRISPR, optimal Cas9 target sites, which include the 5′-GN(17)GG-3′ consensus sequence and a 5′-NGG-3′ protospacer adjacent motif (PAM) (Banerjee et al., 2025b; Farboud, 2017; Farboud and Meyer, 2015), are limited in the AT-rich genome of S. stercoralis. Thus, the use of other Cas proteins that favor AT-rich sequences, such as the Cas12 proteins (Yang et al., 2024), would likely enhance genome editing efficiency in S. stercoralis. In addition, the implementation of other CRISPR/Cas systems for activating or repressing transcription, editing RNA sequences, or altering RNA levels (Knott and Doudna, 2018) would further expand the S. stercoralis molecular genetic toolkit. Additional methods for conditional knockdown of gene function, such as the auxin inducible degradation system (Ashley et al., 2021; Zhang et al., 2015), would also be useful, especially in cases where gene knockout is lethal. Third, bipartite expression systems such as the Cre/Lox (Flavell et al., 2013), Q (Monsalve et al., 2019) and Split cGAL (Wang et al., 2017) systems would be invaluable for precise, inducible genetic targeting of specific cells or subsets of cells. These approaches should be easily adaptable to S. stercoralis using current transgenesis or CRISPR editing methods. Fourth, the development of additional transcriptional reporters that can be used as markers for CRISPR mutagenesis would make it straightforward to generate double and triple knockout lines by introducing multiple mutations sequentially. Fifth, the identification of S. stercoralis genomic safe harbor sites would enable the specific, targeted integration of transgenes using CRISPR without disruption to neighboring genes or uncontrolled copy number; this approach is routinely used for transgene integration in C. elegans (El Mouridi et al., 2022; Malaiwong et al., 2023; Yoshina et al., 2016). Finally, as an increasing number of stable transgenic and knockout lines of S. stercoralis are generated, an efficient cryopreservation protocol will be essential, given that maintenance of S. stercoralis stable lines in gerbils is expensive and labor-intensive. Cryopreservation protocols are already available for some parasitic nematodes, including hookworms (Li et al., 2023; Vacca et al., 2024), and may be adaptable to S. stercoralis. The addition of these methods to the S. stercoralis toolkit would greatly facilitate studies of the molecular and cellular mechanisms that enable parasitic nematodes to find, invade, and infect human hosts.

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Footnotes

Edited by: Mostafa Zamanian, University of Wisconsin-Madison, United States

Reviewed by: Paul J. Brindley, George Washington University, United States