Astacin metalloproteases in human-parasitic nematodes

Abstract

Parasitic nematodes infect over 2 billion individuals worldwide, primarily in low-resource areas, and are responsible for several chronic and potentially deadly diseases. Throughout their life cycle, these parasites are thought to use astacin metalloproteases, a subfamily of zinc-containing metalloendopeptidases, for processes such as skin penetration, molting, and tissue migration. Here, we review the known functions of astacins in human-infective, soil-transmitted parasitic nematodes – including the hookworms Necator americanus and Ancylostoma duodenale, the threadworm Strongyloides stercoralis, the giant roundworm Ascaris lumbricoides, and the whipworm Trichuris trichiura – as well as the human-infective, vector-borne filarial nematodes Wuchereria bancrofti, Onchocerca volvulus, and Brugia malayi. We also review astacin function in parasitic nematodes that infect other mammalian hosts and discuss the potential of astacins as anthelmintic drug targets. Finally, we highlight the molecular and genetic tools that are now available for further exploration of astacin function and discuss how a better understanding of astacin function in human-parasitic nematodes could lead to new avenues for nematode control and drug therapies.

Introduction

Parasitic nematodes infect over 2 billion people globally and are responsible for chronic, debilitating diseases – primarily within disadvantaged communities (Buonfrate et al., 2020; Loukas et al., 2021; Riaz et al., 2020). Individuals living in tropical and subtropical areas that experience poor sanitation, lower socioeconomic status, and inadequate access to healthcare are disproportionally impacted by parasitic nematodes (Beknazarova et al., 2016; Loukas et al., 2021; Magalhaes et al., 2023; McKenna et al., 2017). Human-parasitic nematodes differ in their mode of transmission – some infect through skin penetration, some infect by passive ingestion when they are swallowed, and some are vector-borne (Wakelin, 1996). The hookworms Necator americanus and Ancylostoma duodenale, as well as the threadworm Strongyloides stercoralis, are gastrointestinal nematodes that infect primarily by skin penetration (Colella et al., 2021; Hotez et al., 2004; Page et al., 2018; Schad, 1989; Viney, 2006; Viney, 2017; Viney and Lok, 2015). The giant roundworm Ascaris lumbricoides and the whipworm Trichuris trichiura are also gastrointestinal nematodes, but they infect via passive ingestion (Ahmed, 2023; Dold and Holland, 2011). Filarial nematodes are vector-borne and include Onchocerca volvulus, the causative agent of river blindness; Wuchereria bancrofti and Brugia malayi, which cause lymphatic filariasis; and the African eye worm Loa loa, which causes Loa loa filariasis (Cross, 1996; Hotez et al., 2008; Padgett and Jacobsen, 2008).

Gastrointestinal nematode infections cause malnutritional, digestive, developmental, and inflammatory problems that can result in severe morbidity (Ahmed, 2023; Czeresnia and Weiss, 2022; Dold and Holland, 2011; Loukas et al., 2021; Wang et al., 2008). In addition, S. stercoralis can cause disseminated infections in immunosuppressed individuals that are often fatal (Buonfrate et al., 2023; Czeresnia and Weiss, 2022; Loukas et al., 2021; Mejia and Nutman, 2012). In contrast to gastrointestinal nematodes, filarial nematodes are transmitted through insect intermediate vectors and cause chronic, tissue-damaging infections that are often disfiguring and disabling (Babu and Nutman, 2012; Cross, 1996; Hoerauf, 2008; Hotez et al., 2008; Nutman, 2013; Taylor et al., 2010).

The aforementioned nematode infections are classified as Neglected Tropical Diseases (NTD), which emphasizes the major health and economic burdens these diseases cause globally (Hotez and Lo, 2020). Efforts to control nematode infections are impaired by the need for frequent and repeated anthelmintic drug treatments (Fissiha and Kinde, 2021; Jia et al., 2012). Mass drug administration has reduced disease burden in some endemic areas; however, due to the reliance on a limited number of anthelmintic drugs, including ivermectin and albendazole, it is considered likely that drug-resistant parasite strains will develop (Pilotte et al., 2022). Suboptimal drug efficacy against human-parasitic nematodes has been reported for several of the frontline anthelminthics, challenging scientists to develop novel ways to detect and treat these infections (De Clercq et al., 1997; Herath et al., 2022; Moreno et al., 2021; Mutombo et al., 2019; Repetto et al., 2018).

Many human-parasitic nematodes have complex life cycles, which further complicates disease control efforts (Figure 1). In the case of S. stercoralis, the parasites infect hosts as developmentally arrested infective third-stage larvae (iL3s). The iL3s are soil-dwelling and invade hosts by penetrating through exposed skin. After entering a host, the iL3s resume development and migrate through the host, ultimately ending up as parthenogenetic female adults in the small intestine (Schad et al., 1989). The female adults produce progeny that follow one of three developmental paths: 1) they can exit the host and develop in the soil into iL3s; 2) they can exit the host and develop in the soil into free-living male and female adults, which produce progeny that develop into iL3s; or 3) they can develop into autoinfective larvae (L3a) inside the host and reinfect the same host in a process called autoinfection (Page et al., 2018; Schad, 1989; Schad et al., 1989; Viney, 2006; Viney, 2017; Viney and Lok, 2015). Hookworms are like S. stercoralis in that soil-dwelling iL3s penetrate host skin, but they cannot develop into free-living adults or undergo autoinfection (Colella et al., 2021; Ghodeif and Jain, 2023; Hotez et al., 2004). Ascaris and Trichuris species are orally transmitted when infective (embryonated) eggs are ingested, and both Ascaris and Trichuris adults ultimately reside in the intestine (de Lima Corvino and Horrall, 2023; Dold and Holland, 2011; Viswanath et al., 2023).

Figure 1. The life cycles of selected human-parasitic nematodes, with potential roles of astacins indicated.

Schematics of the life cycles of the skin-penetrating threadworm Strongyloides stercoralis (A), a skin-penetrating hookworm (B), the passively ingested worm Ascaris lumbricoides (C), and a vector-borne filarial worm (D). L1-L4 = 1^st-4^th larval stages; iL3 = infective third larval stage. Life cycle stages that exist inside the host are indicated by the person; life cycle stages that exist in the environment are indicated by the grass and soil; life cycle stages that exist in an insect vector are indicated by the mosquito. The stages of the life cycle where astacins have been demonstrated or hypothesized to function are indicated by the colored symbols, as defined in the key. Life cycle schematics are adapted with permission from Wheeler et al., 2022 (Wheeler et al., 2022) and were generated in BioRender.

The life cycles of filarial nematodes require passage through an insect intermediate host, adding another layer of complexity as compared to other human-infective parasitic nematodes (Figure 1). Third-stage larvae (L3s) infect humans when they are released into host skin during a blood meal by the insect vector. The larval nematodes then migrate throughout the peripheral circulation and lymphatic system or other organs, depending on the species. The adult filariae produce microfilariae that travel to the skin, blood, or lymphatic system. These microfilariae are then ingested by the insect vector during a blood meal. The microfilariae develop and migrate inside the insect body, ultimately residing as L3s in the insect proboscis, where they are transmitted to a human host when the insect takes another blood meal (Brattig, 2004; Gyasi et al., 2023; Newman and Juergens, 2023; Paily et al., 2009).

Despite their life cycle differences, all nematodes, both parasitic and non-parasitic, follow a similar developmental progression in which the worm develops through four larval stages before reaching the adult stage (Page et al., 2014). Each developmental stage transition involves a molt, where the nematode sheds its cuticle and replaces it with a newly synthesized cuticle. The first cuticle is synthesized during late embryogenesis and subsequent cuticles are synthesized during each larval stage, prior to the molt. Thus, a new cuticle is synthesized five times during nematode development (Page and Johnstone, 2007; Page et al., 2014).

Nematode astacin proteins comprise a large group of zinc-dependent metalloproteases that are thought to play critical roles during multiple steps of the nematode life cycle, including embryonic development and molting. In addition, parasitic nematodes likely employ astacins for an expanded repertoire of functions, including penetration of host skin, migration through host tissue, and digestion of host tissue (Figure 1). In a few cases, a requirement for a specific parasitic nematode astacin has been demonstrated. In other cases, a requirement for one or more astacins has been suggested using chemical inhibitors. Here, we review these studies and discuss their implications for nematode control and the development of new anthelmintic drugs. We also discuss how the recent expansion of the human-parasitic nematode genetic toolkit will facilitate future functional studies of nematode astacins.

Astacins comprise a subfamily of metalloproteases

Human-parasitic nematodes utilize myriad proteolytic enzymes to penetrate host skin, migrate within host tissue, molt, digest host tissue, and establish an infection inside their host (Abuzeid et al., 2020; Caffrey et al., 2018; Joshi and Mishra, 2022; Knox, 2011; Yang et al., 2015). Classes of proteolytic enzymes include aspartic, glutamic, cysteine, serine, and threonine proteases as well as metalloproteases (Lopez-Otin and Bond, 2008). Metalloproteases, which require a metal for their catalytic action, are the most diverse of the protease classes and consist of over 70 different families (Rawlings et al., 2018). The astacins comprise a subfamily of metalloproteases that are found in many animal species, including those within the phylum Nematoda (Gomis-Rüth and Stöcker, 2023). Metalloproteases are designated as astacins based on the sequence and structure of their catalytic domain (Gomis-Rüth and Stöcker, 2023; Gomis-Rüth et al., 2012). The term ‘astacin’ is derived from the European crayfish Astacus astacus, where the enzyme was first isolated from the crustacean’s digestive secretions (Sonneborn et al., 1969). The astacin proteins belong to the metzincin superfamily of zinc-dependent proteases (named for their conserved Met residue in the active site and catalytic zinc ion); other members of the superfamily include the matrix metalloproteases (MMP), serralysins, and adamalysins (Gomis-Rüth and Stöcker, 2023). Astacins and adamalysins are categorized into two subfamilies (M12A and M12B, respectively) in the zinc metalloprotease M12 family by the MEROPS database of peptidases, while MMPs are part of the M10 family of metzincins (Rawlings et al., 2018).

All astacin proteins contain a signal peptide at the N-terminus, followed by a prodomain, followed by the zinc-binding catalytic domain (Gomis-Rüth and Stöcker, 2023) (Figure 2A). Many astacins also contain additional domains, which are primarily found downstream of the catalytic domain. For example, some nematode astacins also contain EGF (epidermal growth factor-like), CUB (complement C1r/C1s, Uegf, BMP1), and/or TSP (thrombospondin-like) domains (Gomis-Rüth and Stöcker, 2023) (Figure 2A). Astacins are produced as inactive zymogens that are activated by cleavage of the prodomain, and they can be either membrane-bound or secreted by cells (Gomis-Rüth and Stöcker, 2023; Gomis-Rüth et al., 2012; Möhrlen et al., 2001; Sterchi et al., 2008).

Figure 2. The structure of astacin proteins.

A. Schematic of the functional domains found in three astacin proteins: A. astacus astacin, S. stercoralis strongylastacin, and C. elegans HCH-1. All astacins contain an N-terminal signal peptide (S), a prodomain (PD), and a zinc-dependent catalytic peptidase domain (CPD). Many astacins also contain additional domains, including epidermal growth factor-like (EGF); complement C1r/C1s, Uegf, Bmp1 (CUB); and threonine-rich (TRD) domains. Domains are shown ordered in the 5' to 3' direction; domain sizes are approximate and inter-domain intervals are not to scale. Figure is adapted with permission from Gomez Gallego et al., 2005 (Gomez Gallego et al., 2005). B. Protein alignment of S. stercoralis strongylastacin and A. astacus astacin. Alignment was generated using UniProt. Domains were identified using Prosite and InterPro; motifs were identified based on motif descriptions in Gomis-Rüth and Stöcker, 2023 (Gomis-Rüth and Stöcker, 2023). C. Structure of the predicted peptidase domain of S. stercoralis strongylastacin. The His and Glu residues that bind to the zinc ion are in blue; selected residues of the methionine turn are in magenta. The N- and C-terminal regions of astacin are depicted by yellow arrows. Figure is reprinted with permission from Gomez Gallego et al., 2005 (Gomez Gallego et al., 2005).

Based on structural studies of the A. astacus astacin, the prototypical member of the astacin subfamily, the general structure of the catalytic domain consists of two subdomains separated by a deep active-site cleft that contains zinc at the bottom of the cleft (Bode et al., 1992; Gomis-Rüth and Stöcker, 2023). The active-site cleft contains a zinc-binding HEXXHXXGXXH consensus motif (where “X” represents any residue) that consists of three essential zinc-binding histidine (H) residues as well as a water-linked glutamic acid (G) residue that is required for catalysis (Gomis-Rüth and Stöcker, 2023; Yiallouros et al., 2000) (Figure 2B). The active site also contains a downstream tyrosine residue that participates in substrate binding and stabilization during catalysis (Bode et al., 1992; Gomis-Rüth and Stöcker, 2023; Grams et al., 1996; Yiallouros et al., 2000). The tyrosine residue is located two positions after a “met-turn” methionine, which participates in metal-binding. The tyrosine and methionine are found in a conserved astacin SIMHY-motif (Gomis-Rüth and Stöcker, 2023) (Figure 2B). Thus, the zinc ion in the active site is bound to the protein by a penta-coordinated system involving the three histidine residues, a water molecule, and the tyrosine residue (Grams et al., 1996; Stöcker et al., 1993; Stöcker et al., 1995) (Figure 2C). Astacins differ in their ligand specificity due to key sequence differences in the substrate-binding regions of the protein (Stöcker et al., 1993; Stöcker et al., 1995).

Astacin gene families in human-parasitic nematodes

There are six astacin homologs in mammals (astl, bmp1, mepa, mepb, tll1, and tll2), several of which function during the early stages of fertilization, embryogenesis, and inflammation (Gomis-Rüth and Stöcker, 2023; Herzog et al., 2019). Astacin dysfunction has been implicated in conditions such as cancer, fibrosis, inflammatory bowel disease, and nephritis (Gomis-Rüth and Stöcker, 2023; Herzog et al., 2019). Nematode genomes encode vastly expanded astacin gene families in comparison to mammals. The free-living, non-parasitic nematode Caenorhabditis elegans has 40 astacin genes (called nas genes for “nematode astacins”), whereas S. stercoralis has over 200 astacin genes and A. duodenale has over 160 astacin genes (Consortium, 2019; Hunt et al., 2016; Möhrlen et al., 2003; Park et al., 2010; Xu et al., 2019). In C. elegans, the astacin genes tend to be expressed in tissues exposed to the external environment, such as the pharynx, intestine, and hypodermis (Park et al., 2010). Their functions include cuticle digestion during molting, eggshell digestion during hatching, tissue remodeling during development, and the digestion of food (Möhrlen et al., 2003; Park et al., 2010).

A broad survey of nematode genomes revealed greatly expanded astacin families in skin-penetrating nematodes, including both Strongyloides species and hookworms (Consortium, 2019; Hunt et al., 2016; Hunt et al., 2017; Martin-Galiano and Sotillo, 2022) (Figure 3). In the Strongyloides lineage, the expansion of the astacin gene family correlates with the evolution of parasitism (Consortium, 2019; Hunt et al., 2016; Hunt et al., 2017; Martin-Galiano and Sotillo, 2022). This argument is supported by comparison to the closest free-living, non-parasitic relative of Strongyloides, Rhabditophanes diutinus, which has only 34 astacin genes (Consortium, 2019; Hunt et al., 2016; Martin-Galiano and Sotillo, 2022). Thus, astacins have been proposed to have critical functions during parasite-specific processes such as host invasion and host tissue migration, which require digestion of host skin or other tissues, in addition to processes such as molting and tissue remodeling that are conserved between parasitic and non-parasitic nematodes (Hunt et al., 2016; Hunt et al., 2017; Martin-Galiano and Sotillo, 2022) (Figure 3).

Figure 3. Expansion of the astacin family in skin-penetrating nematodes.

Phylogeny of astacin proteins in the phyla Nematoda (including free-living and parasitic nematodes in Clades I–V) and Platyhelminthes (including cestodes and trematodes). Parasitic nematodes in Clade IV (which includes Strongyloides species) and Clade V (which includes hookworms) have greatly expanded astacin families. Selected astacins that have been functionally studied are labeled. S. stercoralis strongylastacin is thought to contribute to skin penetration (Gomez Gallego et al., 2005; McClure et al., 2023; McKerrow et al., 1990), A. caninum MTP-1 is thought to contribute to skin penetration and tissue migration (Hawdon et al., 1995; McClure et al., 2023; Williamson et al., 2006; Zhan et al., 2002), B. malayi NAS-36 contributes to molting (Stepek et al., 2011), and O. volvulus AST-1 is thought to contribute to tissue migration (Borchert et al., 2007). Figure is reprinted with permission from Martín-Galiano and Sotillo, 2022 (Martin-Galiano and Sotillo, 2022).

In contrast to skin-penetrating nematodes, filarial and passively ingested nematodes have relatively small astacin gene families. The filarial nematodes B. malayi, O. volvulus, and W. bancrofti each have 17 astacin genes (Consortium, 2019; Martin-Galiano and Sotillo, 2022). Similarly, A. lumbricoides has 33 astacin genes, while T. trichiura has 14 (Consortium, 2019; Martin-Galiano and Sotillo, 2022). Some of the astacin genes in these species are most closely related to C. elegans astacins involved in embryonic development and molting; these genes appear to have similar functions in the parasites (Borchert et al., 2007; France et al., 2015; Martin-Galiano and Sotillo, 2022; Ren et al., 2021a; Stepek et al., 2011). However, other astacins in these species are thought to facilitate tissue migration (Borchert et al., 2007; Ren et al., 2021b).

High-throughput transcriptomic and proteomic analyses of parasitic nematodes support an important role for astacins in nematode parasitism. Many astacin genes show highly enriched expression in infective larvae and/or parasitic adults relative to other life stages (Baskaran et al., 2017; Hunt et al., 2018; Hunt et al., 2016; Hunt et al., 2017; Wang et al., 2010; Yoshida et al., 2012). For example, in S. stercoralis, over half of the astacin genes were found to be upregulated in parasitic adult females relative to free-living adult females (Hunt et al., 2016; Hunt et al., 2017). Astacins are also highly abundant in excretory-secretory (ES) products from parasitic adults and infective larvae (Hewitson et al., 2011; Lun et al., 2003; Ren et al., 2021b; Soblik et al., 2011; Sotillo et al., 2014; Varatharajalu et al., 2011). Thus, astacins are ideally localized to interact with host tissue. Interestingly, astacins may contribute to host specificity (i.e., the restricted host range of certain parasitic nematode species) as an RNA-sequencing analysis of the rat parasite Strongyloides ratti revealed that several astacin genes are differentially expressed between parasitic adults isolated from the small intestines of either their natural host (the rat) or a laboratory host (the gerbil) (Jaleta et al., 2017). In addition, expression of an astacin gene in the rat parasite Nippostrongylus brasiliensis was robustly downregulated upon adaptation to an immunocompromised host (Ferguson et al., 2023). Together, these results suggest that the expression of specific astacins may be stimulated by contact between a parasitic nematode and its desired host and may help the parasite suppress the host immune system.

Although functional characterizations of astacin proteins in human-parasitic nematodes remain limited, studies of human-parasitic nematodes as well as other mammalian-parasitic nematodes have implicated astacins in diverse processes such as embryonic development, molting, exsheathment, skin penetration, tissue migration, and intra-host feeding. Thus, parasitic nematode astacins are likely to be essential for multiple stages of the parasite life cycle and multiple steps of the parasite-host interaction.

The role of astacins during embryonic development in human-parasitic nematodes

Several astacins have been implicated in embryonic development. C. elegans nas-35 (also called dpy-31) is essential for formation of the embryonic exoskeleton, and loss of nas-35/dpy-31 function is lethal (Novelli et al., 2004). Mutation of the C. elegans hch-1/nas-34 gene results in hatching defects, as mutant larvae are unable to digest the eggshell (Hishida et al., 1996). In addition, RNA interference (RNAi)-mediated knockdown of C. elegans nas-9 yielded low levels of embryonic lethality, suggesting NAS-9 may be required for embryonic development (Maeda et al., 2001; Möhrlen et al., 2003). While astacin function during embryonic development in parasitic nematodes has not yet been studied, the presence of homologous astacin genes and collagen genes, which are essential for proper egg and embryo development, in the parasitic species suggests that some of the parasitic nematode astacin genes may have similar functions during embryonic development (Martin-Galiano and Sotillo, 2022; Page et al., 2014).

The role of astacins during molting in human-parasitic nematodes

Molting, or ecdysis, is the process by which a worm sheds its cuticle and develops a new one. Molting is an essential developmental process for both parasitic and non-parasitic nematodes. All nematodes undergo four larval molts before reaching adulthood (Lažetić and Fay, 2017; Page et al., 2014). Parasitic nematodes molt before and after infecting a host. For example, filarial worms molt twice in the insect intermediate vector and twice in the mammalian host (Cross, 1996; Newman and Juergens, 2023). In contrast, hookworms molt twice outside the host and twice inside the host (Ghodeif and Jain, 2023) (Figure 1). In the case of hookworms, larvae also undergo a molt-like process called exsheathment – the iL3s retain the second-stage larval cuticle as a loose outer sheath and then exsheath shortly before or after host entry (Loukas et al., 2021). Studies of skin penetration in N. americanus found iL3 sheaths on the skin surface, suggesting that N. americanus iL3s exsheath immediately before or during skin penetration (Goodey, 1922).

Nematode cuticles are rich in collagen, and collagen-degrading astacins mediate cuticle degradation during molting (Page et al., 2014). In C. elegans, mutation or RNAi-mediated knockdown of nas-37 results in incomplete ecdysis, such that the old cuticle is only partially shed during molting (Davis et al., 2004; Kamath et al., 2003; Maeda et al., 2001; Möhrlen et al., 2003; Suzuki et al., 2004). The nas-37 gene is expressed in the hypodermis shortly before each molt, and NAS-37 protein accumulates at the anterior end of the worm, where it functions to degrade the old cuticle (Davis et al., 2004). RNAi-mediated knockdown of the nas-36 gene caused a similar, although less severe, defect (Suzuki et al., 2004). Thus, both nas-36 and nas-37 are required for normal molting in C. elegans. Several other astacin genes are required for molting in C. elegans, including nas-35/dpy-31 (Kamath et al., 2003; Maeda et al., 2001;Novelli et al., 2004). While null mutations of nas-35/dpy-31 are lethal, hypomorphic mutations in the gene result in a “dumpy” (Dpy) phenotype characterized by a “short and fat” body shape, indicative of a defect in cuticle formation (Novelli et al., 2004).

The role of astacins during molting in parasitic nematodes is less well understood, largely because the molecular toolkit for studying gene function in most parasitic nematodes remains limited. Early studies found that a zinc-containing metalloprotease present in the exsheathing fluids of the ruminant parasite Haemonchus contortus induces the formation of a refractile ring near the apical end of the cuticle; this ultimately leads to removal of the cuticular cap, which provides a hole through which the larva rapidly exits the sheath (Gamble et al., 1989a; Gamble et al., 1989b). Purified C. elegans NAS-37 protein applied to isolated H. contortus cuticles was found to induce refractile ring formation, raising the possibility that related astacins in H. contortus may be involved in exsheathment (Davis et al., 2004). Subsequently, a homolog of C. elegans nas-36 was identified in both H. contortus and B. malayi (Stepek et al., 2011). Expression of B. malayi nas-36 in the C. elegans nas-36 or nas-37 mutant background was sufficient to rescue the molting defect, suggesting that B. malayi nas-36 is a functional metalloprotease that may function during B. malayi molting and exsheathment (Stepek et al., 2011). RNAi-mediated knockdown of the astacin gene nas-33 in H. contortus resulted in a molting defect in L1 larvae that was ultimately lethal, thus directly implicating nas-33 in parasite molting (Huang et al., 2021) (Figure 4). Interestingly, knockdown of C. elegans nas-33 did not result in a molting phenotype, suggesting possible functional divergence or species-specific differences in redundancy (Huang et al., 2021).

Figure 4. RNAi-mediated knockdown of the H. contortus gene nas-33 causes molting defects.

A. Knockdown of nas-33 in H. contortus first-stage larvae in an RNAi feeding assay resulted in a molting defect in which the old cuticle remained partly attached to the second-stage larva after the molt (Huang et al., 2021). B-D. Enlarged images of the indicated regions in A; sites where the old cuticle remained attached to the second-stage larva are indicated with white arrows (Huang et al., 2021). Images are reprinted from Huang et al., 2021 (Huang et al., 2021).

NAS-35/DPY-31 homologs have been identified in parasitic nematodes, including B. malayi, H. contortus, the ruminant parasite Teladorsagia circumcincta, and the food-borne pork worm Trichinella spiralis (France et al., 2015; Ren et al., 2021a; Stepek et al., 2010; Stepek et al., 2015). H. contortus nas-35/dpy-31 was able to rescue the C. elegans nas-35/dpy-31 phenotype, and both H. contortus and B. malayi NAS-35/DPY-31 were able to digest the C. elegans cuticle collagen SQT-1 (Stepek et al., 2010). Moreover, RNAi-mediated knockdown of T. spiralis NAS-35/DPY-31 resulted in cuticle defects and reduced worm burden in infected mice (Ren et al., 2021a). Together, these studies suggest broad conservation of NAS-35/DPY-31 function across species. However, the contribution of astacins to molting in S. stercoralis, hookworms, and many other human-parasitic nematodes remains to be investigated.

The role of astacins during skin penetration by skin-penetrating parasitic nematodes

Both hookworms in the genera Necator and Ancylostoma, and threadworms in the genus Strongyloides, infect hosts by penetrating through host skin (McClure et al., 2023). Although skin-penetration behavior has yet to be characterized in detail, iL3s are known to penetrate the skin head-first (Goodey, 1925; Zaman et al., 1980). They enter the skin either by puncturing directly through the outer layer of skin, called the stratum corneum, or by crawling into hair follicles (Abadie, 1963; Goodey, 1925; Matthews, 1972; Vetter and Leegwater-vd Linden, 1977; Zaman et al., 1980). The process of burrowing into the skin involves digestion of the skin by ES products secreted by iL3s (Dresden et al., 1985; Hotez et al., 1990; Matthews, 1982; Smith, 1976).

Biochemical analysis of iL3 secretions from S. stercoralis, N. americanus, A. duodenale, and A. caninum found that they contain metalloproteases capable of digesting the dye-labeled collagen substrate azocoll (Hotez et al., 1990; Matthews, 1982; McKerrow et al., 1990; Williamson et al., 2006). When the ES products of S. stercoralis iL3s were treated with chemical inhibitors that specifically target zinc metalloproteases, the ES products were no longer able to degrade azocoll (McKerrow et al., 1990) (Figure 5A). Moreover, exposure of S. stercoralis iL3s to zinc metalloprotease inhibitors blocked skin penetration in an ex vivo assay, suggesting a requirement for zinc metalloproteases during skin penetration (McKerrow et al., 1990) (Figure 5B). Fractionation of the ES products identified an approximately 40 kDa metalloprotease present at high abundance in iL3 secretions; this protease was initially named Ss40 (McKerrow et al., 1990) but was later identified as an astacin and renamed strongylastacin (Gomez Gallego et al., 2005) (Figure 2). Similarly, an astacin from the dog hookworm Ancylostoma caninum, named Ac-MTP-1, was found to be capable of digesting proteins from connective tissue (Williamson et al., 2006). Incubating A. caninum iL3s with either anti-Ac-MTP-1 serum or small-molecule metalloprotease inhibitors blocked skin penetration in an ex vivo assay with dog skin, suggesting that Ac-MTP-1 may be required for skin penetration (Williamson et al., 2006). More recent analyses of the secretome of skin-penetrating iL3s have revealed that they are highly enriched in astacins (Cuesta-Astroz et al., 2017; Maeda et al., 2019; Soblik et al., 2011). Taken together, these results suggest that astacins are essential for skin-penetrating iL3s to penetrate host skin. However, the functions of individual astacin genes during skin penetration remain to be determined.

Figure 5. Metalloprotease inhibitors prevent skin penetration by S. stercoralis iL3s.

A. Incubation of ES products from S. stercoralis iL3s with metalloprotease inhibitors blocks their proteolytic activity, suggesting that protein degradation by iL3s requires metalloproteases. ES products isolated from S. stercoralis iL3s were incubated with either metalloprotease inhibitors (2.5 mM 1,10-phenanthroline; 10 mM HONHCOCH (CH₂CH₂CH Me₂) CO-Ala-Gly-NH₂; or 1 mM HO₂C-CH₂-Phe-Leu), serine protease inhibitors (5 mM PMSF, 100 μg/mL Aprotinin, or 100 μg/mL α-1-proteinase inhibitor), a cysteine protease inhibitor (5 mM NEM), or an aspartic protease inhibitor (50 μg/mL pepstatin). The ES products were then exposed to the substrate azocoll, and the % inhibition of azocoll degradation was determined. Only exposure to metalloprotease inhibitors blocked azocoll degradation. B. Incubation of S. stercoralis iL3s with metalloprotease inhibitors blocks skin penetration in an ex vivo skin penetration assay. iL3s incubated with either metalloprotease inhibitors (2 mM 1,10-phenanthroline; 2 mM HONHCOCH (CH₂CH₂CH Me₂) CO-Ala-Gly-NH₂; or 2 mM HO₂C-CH₂-Phe-Leu) or a serine protease inhibitor (2 mM PMSF) were placed on rat skin, and the % inhibition of skin penetration was determined based on the number of iL3s that penetrated the skin. Only exposure to the metalloprotease inhibitors blocked skin penetration. For A-B, percent inhibition was calculated for each chemical relative to the vehicle control. Data are replotted with permission from McKerrow et al., 1990 (McKerrow et al., 1990).

The role of astacins during host tissue migration and feeding in human-parasitic nematodes

Astacin proteins are highly represented in the ES products of the intra-host life stages of parasitic nematodes; thus, it has been hypothesized that astacins are critical for digestion of host tissue during tissue migration and intra-host feeding. When the developmentally arrested iL3s of skin-penetrating nematodes enter a host, they exit developmental arrest and resume growth in a process called activation (Hawdon et al., 1992; Mendez et al., 2022). The A. caninum astacin Ac-MTP-1 is expressed in the pharynx and is secreted by activated iL3s, suggesting it may contribute to the ability of iL3s to migrate through host tissue (Hawdon et al., 1995; Zhan et al., 2002). A different A. caninum astacin, Ac-MTP-2, is expressed in the pharyngeal glands of parasitic adults and is found in their ES products, also suggestive of a function in digesting host tissue (Feng et al., 2007).

Astacins are also thought to play a role in tissue migration in filarial and passively ingested nematodes. T. spiralis infects humans when encysted larvae in the muscle tissue of an infected host are ingested. These larvae are then activated into intestinal infective larvae by host digestive enzymes inside the stomach and subsequently burrow into the host intestinal epithelium (Furhad and Bokhari, 2023; Ren et al., 2021b). Astacins are present in the ES products of T. spiralis intestinal infective larvae and are therefore well-positioned to mediate tissue migration by the larvae (Ren et al., 2021b). Similarly, in O. volvulus, astacins are found in the ES products of the tissue-migrating microfilariae, suggestive of a function during tissue migration through the human host (Borchert et al., 2007; Haffner et al., 1998). Astacins are also present in the predicted secretome of the giant porcine roundworm Ascaris suum and are hypothesized to function during tissue migration (Jex et al., 2011).

Astacins may contribute to nematode feeding inside the host. A zinc metalloprotease in H. contortus is expressed around the time larvae begin to feed inside their host (Gamble et al., 1996), and characterization of the intestinal transcriptomes of N. americanus and A. caninum adults identified astacin genes that are highly expressed in the gut (Ranjit et al., 2006), consistent with a function during feeding. However, as with skin penetration, a direct requirement for individual astacins during tissue migration and feeding on host tissue has yet to be demonstrated.

Implications for nematode control

Astacins have long been of interest as targets for novel anthelmintic drugs. Given that astacins are likely critical for multiple steps of the parasitic nematode life cycle (Figure 1), interruption of astacin function could severely compromise or eliminate the parasite’s ability to propagate and establish an infection. Moreover, it is likely that astacin-targeting anthelmintics would be active against all developmental stages of parasitic nematodes, which is an attractive design element for future chemotherapeutics. Current frontline anthelmintics such as albendazole and ivermectin target β-tubulin and glutamate-gated chloride channels in parasitic nematodes, respectively (Deep et al., 2023). Both compounds have parasite-specific targets, thus reducing off-target hits. However, continued reliance on these drugs has increased the risk of drug resistance and novel therapeutics are urgently needed (Fissiha and Kinde, 2021; Jia et al., 2012; Pilotte et al., 2022).

Anti-astacin therapeutics are an emerging field in drug development, particularly for the treatment of human diseases (Bond, 2019). Inhibitors of the astacins meprin α and β have been investigated for the treatment of a wide array of disease processes, including fibrosis, cancer, and inflammatory bowel disease (Bayly-Jones et al., 2022; Jana et al., 2024; Linnert et al., 2021; Talantikite et al., 2018; Tan et al., 2021; Turtle et al., 2012). Several studies have also investigated the potential of small-molecular inhibitors to block nematode astacin function. One of the astacins that has been examined as a possible drug target is NAS-35/DPY-31. Small-molecule screens for inhibitors of recombinant B. malayi and T. circumcincta NAS-35/DPY-31 identified several inhibitors of the protein (France et al., 2015; Stepek et al., 2015). These include two novel tripeptide hydroxamic acids (France et al., 2015); marimastat, a drug that has been explored for the treatment of cancer and epilepsy (King et al., 2003; Pijet et al., 2020; Renkiewicz et al., 2003; Sparano et al., 2004; Wojtowicz-Praga et al., 1998); and actinonin, a naturally occurring antibiotic (France et al., 2015; Sina et al., 2009; Stepek et al., 2015). Exposure of C. elegans, T. circumcincta, and H. contortus to the tripeptide hydroxamic acids resulted in a Dpy phenotype, indicating that the inhibitors can disrupt molting and induce cuticle defects (France et al., 2015; Stepek et al., 2015). These results raise the possibility that inhibitors of NAS-35/DPY-31 could act as broad-spectrum anthelmintics.

Another approach to nematode control is vaccine development. Astacins have been identified as promising antigens for helminth vaccines due to their location at the parasite-host interface. There are currently no commercially available vaccines against human-parasitic nematodes, although several vaccines are in various stages of development, including in preclinical or clinical trials (Abraham et al., 2021; Adegnika et al., 2021; Chapman et al., 2021; Diemert et al., 2022; Lustigman et al., 2018; Mouwenda et al., 2021; Puchner et al., 2023; Ryan et al., 2021; Ryan et al., 2023; Zhan et al., 2022). Some of these vaccines use the N. americanus aspartic protease Na-ASP-1 as a vaccine antigen, demonstrating the potential efficacy of proteases in vaccine development (Diemert et al., 2022; Puchner et al., 2023). Although astacin-based vaccines have not been tested in humans, several astacins, including Ace-MTP-2 from the human-parasitic hookworm Ancylostoma ceylanicum, strongylastacin from S. stercoralis, and Ov-AST-1 from O. volvulus, are immunogenic (Baska et al., 2013; Borchert et al., 2007; Brindley et al., 1995). Moreover, Ace-MTP-1 from A. ceylanicum showed promise as a vaccine antigen when administered to hamsters (Mendez et al., 2005). Ac-MTP-1 from A. caninum also showed early promise as a vaccine antigen when administered to dogs and hamsters (Hotez et al., 2003). Interestingly, a study of new potential vaccine candidates for prevention of H. contortus infection identified an astacin gene that was differentially expressed in resistant vs. susceptible worms (Palevich et al., 2023). This finding raises the possibility that some astacins may be involved in immune evasion, and drugs that target these astacins could be useful in enhancing vaccine efficacy (Palevich et al., 2023).

Finally, astacins may also be valuable tools in the development of new methods for diagnosing nematode infections. While anthelmintic treatments are essential for controlling nematode infections, proper diagnostic methods are also critical for tracking infection rates and providing targeted therapeutics to areas in need (Mutombo et al., 2019). Parasitic infections can be confirmed through several methods, including ova and parasite fecal tests and antigen/antibody or polymerase chain reaction tests of blood, saliva, or stool (Branda et al., 2006; Buonfrate et al., 2015; Khurana and Sethi, 2017; Mutombo et al., 2019; Needham et al., 1996). Many antigen/antibody tests involve antibodies raised against ES products (Harnett, 2014; Sykes and McCarthy, 2011). Some recombinant helminth proteases have shown promise as diagnostic antigens (Farid, 2023; Sun et al., 2018; Zhang et al., 2016), raising the possibility that recombinant nematode astacins could be used as antigens to detect species-specific antibodies for diagnosing nematode infections in humans.

Conclusions and future directions

Here, we have provided an overview of astacin metalloprotease function in parasitic nematodes. We have described the evidence that parasitic nematode astacins contribute to multiple stages of the parasitic nematode life cycle, including embryonic development, molting, skin penetrating, tissue migration, and intra-host feeding. We have also discussed the potential for these enzymes to be exploited for the development of novel therapeutics, vaccines, and diagnostic methods for treating and preventing nematode infections.

The development of astacin-based nematode control and diagnostic strategies would be greatly aided by a better understanding of the basic biology of these proteins. Until recently, the lack of genetic tools for human-parasitic nematodes precluded in vivo studies of astacin function. Excitingly, the recent development of new tools for the genetic manipulation of some human-parasitic nematodes has enabled more mechanistic studies of parasite genes (Castelletto et al., 2020; Mendez et al., 2022). Notably, techniques for transgenesis and CRISPR-mediated mutagenesis have been developed for S. stercoralis and have led to new insights into many aspects of their basic biology, including chemosensation, thermosensation, locomotion, and development (Adams et al., 2019; Bryant et al., 2018; Bryant et al., 2022; Castelletto et al., 2009; Cheong et al., 2021; Dulovic and Streit, 2019; Gang et al., 2017; Gang et al., 2020; Lok et al., 2017; Wang et al., 2015). Techniques for transgenesis and CRISPR-mediated mutagenesis have also been developed for B. malayi (Higazi et al., 2002; Liu et al., 2020a; Liu et al., 2020b; Liu et al., 2018; Xu et al., 2011), and RNAi has been used successfully in B. malayi and O. volvulus (Castelletto et al., 2020; Kashyap et al., 2019; Luck et al., 2016; Lustigman et al., 2004; Misra et al., 2017; Verma et al., 2017). Leveraging these tools, future studies of astacin function in human-parasitic nematodes should be able to identify the cells that express astacins; identify where and when astacins are secreted; and test the requirement for astacins in various biological processes. A better understanding of astacin function in human-parasitic nematodes will inform future nematode control efforts.