Nematophilic bacteria associated with entomopathogenic nematodes and drug development of their biomolecules.

The importance of Xenorhabdus and Photorhabdus symbionts to their respective Steinernema and Heterorhabditis nematode hosts is that they not only contribute to their entomopathogenicity but also to their fecundity through the production of small molecules. Thus, this mini-review gives a brief introductory overview of these nematophilic bacteria. Specifically, their type species, nematode hosts, and geographic region of isolations are tabulated. The use of nucleotide sequence-based techniques for their species delineation and how pangenomes can improve this are highlighted. Using the Steinernema-Xenorhabdus association as an example, the bacterium-nematode lifecycle is visualized with an emphasis on the role of bacterial biomolecules. Those currently in drug development are discussed, and two potential antimalarial lead compounds are highlighted. Thus, this mini-review tabulates forty-eight significant nematophilic bacteria and visualizes the ecological importance of their biomolecules. It further discusses three of these biomolecules that are currently in drug development. Through it, one is introduced to Xenorhabdus and Photorhabdus bacteria, their natural production of biomolecules in the nematode-bacterium lifecycle, and how these molecules are useful in developing novel therapies.

Introduction

Nematophilic “nematode loving” bacteria are prokaryotes that are symbiotically associated with members of phylum Nematoda. Three such genera are Xenorhabdus, Photorhabdus, and Serratia, which are symbionts of Steinernematidae, Heterorhabditidae, and Rhabditidae members, respectively. Each of these three families of the order Rhabditida contain entomopathogenic species—all members of Steinernema and Heterorhabditis, all members of the Insectivora—group of Oscheius, and Caenorhabditis briggsae. However, it is their Xenorhabdus, Photorhabdus, or Serratia symbionts that contribute in large part to this trait through both septicemia and toxemia (Bowen et al., 1998; Ffrench-Constant and Waterfield, 2005; Abebe et al., 2011; Clarke, 2020). Hence, Xenorhabdus, Photorhabdus, and a few Serratia strains are also termed entomopathogenic bacteria. Whereas Serratia symbionts form associations with Oscheius and Caenorhabditis hosts, Xenorhabdus and Photorhabdus are more genus-specific associating only with Steinernema and Heterorhabditis hosts, respectively (Table 1).

TABLE 1

Nematophilic bacteria associated with entomopathogenic nematodes (EPNs).

Species	Nematode host of isolation	Geographic origin of nematode	Example of a bioactive molecule produced by the type strain
Xenorhabdus beddingii (Akhurst and Boemare, 1988)	Steinernema longicaudum	Tasmania, Australia (Akhurst, 1983)	Xefoampeptide (Tobias et al., 2017; Kegler and Bode, 2020)
X. bovienii (Akhurst and Boemare, 1988)	S. affine	Tasmania, Australia (Akhurst, 1983)	Xenocyloin (Proschak et al., 2014)
	S. intermedium
	S. kraussei
	S. feltiae
X. budapestensis (Lengyel et al., 2005)	S. bicornutum	Szabadka, Serbia (Tallosi et al., 1995)	Bicornutin (Böszörményi et al., 2009)
X. cabanillasii (Tailliez et al., 2006)	S. riobrave	Weslaco, USA (Cabanillas et al., 1994)	Rhabdopeptide (Reimer et al., 2013)
X. doucetiae (Tailliez et al., 2006)	S. diaprepesi	Martinique, Caribbean (Fischer-Le Saux et al., 1998)	Xenorhabdin (Bode et al., 2015)
X. eapokensis (Kämpfer et al., 2017)	S. eapokense	Eapok, Vietnam (Phan et al., 2006)	GameXPeptide (Tobias et al., 2017; Shi et al., 2022)
X. ehlersii (Lengyel et al., 2005)	S. serratum	Shangdong, China (Qiu et al., 2004)	GameXPeptide (Tobias et al., 2017; Shi et al., 2022)
X. griffiniae (Tailliez et al., 2006)	S. hermaphroditum	Kamal, Indonesia (Stock et al., 2004)
X. hominickii (Tailliez et al., 2006)	S. karii	Kirinyaga, Kenya (Waturu et al., 1997)	Fabclavine (Wenski et al., 2020)
	S. monticolum
X. indica (Somvanshi et al., 2006)	S. thermophilum	New Delhi, India (Sudershan and Singh, 2000)	Taxlllaid (Kronenwerth et al., 2014)
X. innexi (Lengyel et al., 2005)	S. scapterisci	Rivera, Uruguay (Nguyen and Smart, 1990)	Rhabdopeptide/xenortide-like peptide (Zhao L. et al., 2018)
X. japonica (Nishimura et al., 1994)	S. kushidai	Hamakita, Japan (Nishimura et al., 1994)	Lipocitide (Shi et al., 2022)
X. ishibashii (Kuwata et al., 2013)	S. aciari	Haimen, China (Qiu et al., 2005)	Xenorhabdin (McInerney et al., 1991; Bode et al., 2015; Tobias et al., 2017)
X. lircayensis (Castaneda-Alvarez et al., 2021)	S. unicornum	Altos de Lircay, Chile (Castaneda-Alvarez et al., 2021)
X. khoisanae (Ferreira et al., 2013)	S. khoisanae	Villiersdorp, South Africa (Malan et al., 2006)
X. koppenhoeferi (Tailliez et al., 2006)	S. scarabaei	New Jersey, USA (Stock and Koppenhöfer, 2003)
X. kozodoii (Tailliez et al., 2006)	S. arenarium	Voronezh, Russia (Artyukhovsky, 1997)	Xenocoumacin (Park et al., 2009; Tobias et al., 2017)
	S. apuliae
X. magdalenensis (Tailliez et al., 2012)	S. australe	Isla Magdalena, Chile (Edgington et al., 2009)
X. mauleonii (Tailliez et al., 2006)	Steinernema sp.	St. Vincent, Caribbean (Fischer-Le Saux et al., 1998)	Xenoamicin (Zhou et al., 2013)
X. miraniensis (Tailliez et al., 2006)	Steinernema sp.	Mirani, Australia (Akhurst and Boemare, 1988)	Ambactin (Schimming et al., 2014)
X. nematophila (Akhurst and Boemare, 1988)	S. carpocapsae	Virginia, USA (Poinar et al., 1972)	Rhabduscin (Eugenia Nuñez-Valdez et al., 2019)
X. poinarii (Akhurst and Boemare, 1988)	S. glaseri	North Carolina, USA (Poinar, 1978)
	S. cubanum
X. romanii (Tailliez et al., 2006)	S. puertoricense	Puerto Rico, USA (Román and Figueroa, 1994)
X. stockiae (Tailliez et al., 2006)	S. siamkayai	Lohmsak, Thailand (Stock, 1998)	GameXPeptide (Tobias et al., 2017; Shi et al., 2022)
X. szentirmaii (Lengyel et al., 2005)	S. rarum	Cordoba, Argentina (Aguera de Doucet, 1986)	Szentiamide (Ohlendorf et al., 2011)
X. thuongxuanensis (Kämpfer et al., 2017)	S. sangi	Thuongxuan, Vietnam (Phan et al., 2001)	GameXPeptide (Tobias et al., 2017; Shi et al., 2022)
X. vietnamensis (Tailliez et al., 2010)	S. sangi	Xuanmy, Vietnam (Phan et al., 2001)	Benzobactin A (Shi et al., 2022)
Serratia nematodiphila (Zhang et al., 2009)	Oscheius chongmingensis	Chongming Islands, China (Zhang et al., 2009)
S. marcescens (Torres-Barragan et al., 2011)	O. carolinensis	Raleigh, USA (Ye et al., 2010)
	O. safricana	Northwest Province, South Africa (Serepa-Dlamini and Gray, 2018)
Serratia sp. strain TEL (Lephoto Tiisetso et al., 2015)	O. basothovii	Suikerbosrand Nature Reserve, South Africa (Lephoto Tiisetso et al., 2015)
Serratia sp. strain N19 (Zhou et al., 2017)	O. microvilli	Chongming Island, China (Zhou et al., 2017)
Serratia sp. strain SCBI (Abebe et al., 2011)	Caenorhabditis briggsae	Mpumalanga Province, South Africa (Abebe et al., 2010)
Photorhabdus aegyptia (Machado et al., 2021a)	Heterorhabditis bacteriophora	Egypt (Hussein and El-Souud, 2006)	Piscibactin (Shi et al., 2022)
P. akhurstii (Machado et al., 2018)	H. indica	Grande Terre, Guadeloupe Islands (Fischer-Le Saux et al., 1998)
P. asymbiotica (Fischer-Le Saux et al., 1999)	Unknown	San Antonio, USA (Farmer et al., 1989)
P. australis (Machado et al., 2018)	H. gerrardi	Victoria, Australia (Peel et al., 1999; Plichta et al., 2009)	Glidobactin (Tobias et al., 2017)
P. bodei (Machado et al., 2018)	H. beicherriana	Liaoning Province, China (Machado et al., 2018)	Photoxenobactin (Shi et al., 2022)
P. caribbeanensis (Machado et al., 2018)	H. bacteriophora	Basse Terre, Guadeloupe Islands (Fischer-Le Saux et al., 1998)
P. cinerea (Machado et al., 2018)	H. downesi	Ásotthalom, Hungary (Tóth and Lakatos, 2008)
P. hainanensis (Machado et al., 2018)	Heterorhabditis sp.	Hainan Island, China (Akhurst, 1987)
P. heterorhabditis (Ferreira et al., 2014)	H. zealandica	Brits, South Africa (Mothupi, 2016)
P. hindustanensis (Machado et al., 2021b)	Heterorhabditis sp.	Meghalaya, India (Ganguly et al., 2010)
P. kleinii (Machado et al., 2018)	H. georgiana	Ohio, USA (An and Grewal, 2011)
P. kayaii (Machado et al., 2018)	H. bacteriophora	Aksaray, Turkey (Hazir et al., 2003)
P. khanii (Machado et al., 2018)	H. bacteriophora	Clayton, USA (Khan et al., 1976)
P. laumondii (Machado et al., 2018)	H. bacteriophora	Trindad, Trindad and Tobago (Fischer-Le Saux et al., 1998)	Makes Caterpillar Floppy toxin (Daborn et al., 2002)
P. luminescens (Boemare et al., 1993)	H. bacteriophora	Brecon, Australia (Thomas and Poinar, 1979)	3,5-dihydroxy-4-isopropylstilbene (Hu et al., 1997)
P. namnaonensis (Machado et al., 2018)	H. baujardi	Nam Nao, Thailand (Glaeser et al., 2017)	3-isopropyl-4-oxo-2-oxetanecarboxylic acid (Shi et al., 2022)
P. noenieputensis (Machado et al., 2018)	H. noenieputensis	Nelspruit, South Africa (Malan et al., 2011)
P. stackebrandtii (Machado et al., 2018)	H. bacteriophora	Atwood, USA (Grewal et al., 2002)
P. tasmaniensis (Machado et al., 2018)	H. zealandica	Nicholls Rivulet, Australia (Akhurst, 1987)
P. temperata (Fischer-Le Saux et al., 1999)	H. megidis	Nachodka, Russia (Akhurst, 1987)
P. thracensis (Machado et al., 2018)	H. bacteriaphora	Kirklareli, Turkey (Hazir et al., 2003)	GameXPeptide (Tobias et al., 2017; Shi et al., 2022)

Apart from Xenorhabdus and Serratia, other entomopathogenic bacterial symbionts such as Pseudomonas sp. (Ogier et al., 2020) and Alcaligenes sp. (Shan et al., 2019) associate with Steinernema and Oscheius nematodes, respectively.

This classification of Serratia as nematophilic bacteria has caveats. Not all species in the Serratia genus are nematode symbionts (Grimont and Grimont, 2006), and only a few Oscheius-Serratia/Caenorhabditis-Serratia associations are known (Table 1). Moreover, for some of these associations, Serratia were only facultative symbionts—Serratia sp. strain SCBI and Serratia marcescens from C. briggsae and Oscheius carolinensis, respectively (Abebe et al., 2011; Torres-Barragan et al., 2011). Conversely, except one, all characterized Xenorhabdus and Photorhabdus species are natural nematode intestinal symbionts (Table 1). Thus, this mini-review focused on Xenorhabdus and Photorhabdus as nematophilic bacteria.

Xenorhabdus and Photorhabdus bacteria

Xenorhabdus and Photorhabdus are both gram-negative, rod-shaped, peritrichously flagellated, facultative anaerobes of the family Morganellaceae, order Enterobacterales, and class Gammaproteobacteria (Adeolu et al., 2016). They are uniquely characterized by not only having primary and secondary variants but also an endosymbiosis with entomopathogenic nematodes (EPNs). Other distinguishing traits include Photorhabdus as the only terrestrial bioluminescent bacterium genus and Xenorhabdus as the only member of Enterobacterales that does not produce catalase (Boemare and Akhurst, 2006). Despite this taxonomic relatedness, the similar ecological niche of the two is more due to convergent evolution (Chaston et al., 2011). Twenty-seven Xenorhabdus species that were isolated from twenty-seven steinernematids have been described to date (Table 1). However, 100 Steinernema species have been characterized (Bhat et al., 2020) highlighting that at most—because of species with more than one nematode host (Table 1)—63 novel Xenorhabdus species could be added to the genus from these respective under-investigated yet described steinernematids. This prediction can be mathematically supported by determining whether the Xenorhabdus pangenome is open (Medini et al., 2020).

Nucleotide sequence-based techniques are not only the gold standard for prokaryotic species delineation (Chun et al., 2018) but are also useful for either identification of new isolates or emendation of already described taxon. For example, Xenorhabdus sp. strain BMMCB was described as an Xenorhabdus. griffiniae species (Mothupi et al., 2015), but we (Awori et al., 2017) demonstrated that its nucleotide identities values for the recombinase A (recA), phosphoserine transferase (serC), and small subunit ribosomal ribonucleic acid (rRNA) (SSU) genes, with those of the type species, were below the accepted threshold for conspecific strains—97% for protein-coding genes (Tailliez et al., 2010) and 98.7% for SSU gene (Kim et al., 2014).

Two powerful nucleotide sequence-based techniques are average nucleotide identities (ANI) and digital DNA–DNA hybridization (dDDH), which both delineate species by calculating how related two genomes are. The thresholds for conspecific strains are >95% (Richter and Rosselló-Móra, 2009) and >70% (Auch et al., 2010) for ANI and dDDH, respectively. Both were used to reclassify Photorhabdus species (Machado et al., 2018). However, strains S8-52, S9-53, and S10-54 identified as Photorhabdus kleinii had ANI values of 96.7% with the Photorhabdus bodei type strain and Photorhabdus temperata Meg1 had ANI values of 96.3% with the Photorhabdus thracensis type strain demonstrating the difficulty in delineating species of Photorhabdus (Fischer-Le Saux et al., 1998; Tailliez et al., 2010; Machado et al., 2021b) even with these nucleotide-based thresholds (Bobay, 2020). Thus, the use of pangenome analysis for species delineations—as was done in the Prochlorococcus genus (Moldovan and Gelfand, 2018)—is recommended for Photorhabdus systematics when sufficient genome sequences—at least five per species (Medini et al., 2020)—are available.

The nematode-bacterium lifecycle and bacterial biomolecules

The nematode-bacterium lifecycle begins with soil-dwelling infective third larval stage juvenile nematode (IJ3) preying on an insect (Figure 1). Anatomically, IJ3 are third larval stage juvenile nematodes (J3) with a retained second larval stage cuticle that seals both mouth and anus rendering the nematodes into a non-feeding, developmentally arrested, and perennation-like stage (Poinar and Leutenegger, 1968). Steinernematids IJ3 infect an insect only through natural openings, whereas heterorhabditis can additionally gain entry by piercing into the hemocoel using a bursa (Bedding and Molyneux, 1982). Once within, the IJ3s undergo “recovery” (Clarke, 2020) whereby they shed their second larval stage cuticle and release into the hemocoel, their gut bacterial symbionts. For Steinernema, Xenorhabdus would have been previously localized in a receptacle (Stilwell et al., 2018) at the anterior gut whereas, in Heterorhabditis, Photorhabdus would have previously lined the entire gut (Waterfield et al., 2009). Detection of L-proline concentrations >4.8 mM in insect hemolymph triggers an upregulated bacterial secretion of specialized metabolites of various ecological functions (Crawford et al., 2010).

FIGURE 1

Xenorhabdus-Steinernema lifecycle and selected biomolecules that contribute toward nematode fecundity. Free-living infective third-stage juvenile (IJ3) nematodes seek out insects and gain entry through natural openings such as spiracles, and once within the hemocoel, nematodes exit their non-feeding state and release Xenorhabdus gut symbionts. The bacteria secrete a range of biomolecules (1–13) that increase the fecundity of the nematodes. Nematodes go through complete lifecycles thus increasing their numbers and upon depletion of nutrients each J3 re-associates with a few Xenorhabdus bacteria and exit the insect cadaver as an IJ3. J1, J2, J3, and J4 = first, second, third, and fourth larval stage juvenile nematodes, respectively. Benzylacetone (1), rhabduscin (2), xenocyloin (3), cabanillasin (4), biocornutin (5), xenocoumacin 2 (6), odilorhabdin (7), nematophin (8), xenorhabdin (9), xenocoumacin 1 (10), xenortide (11), rhabdopeptide (12), and rhabdopeptide/xenortide-like peptides (13) were created in biorender.com.

Despite the following grouping of biomolecules from both Xenorhabdus and Photorhabdus according to the similarity of ecological function, their biosynthesis is species-specific. The first grouping is insecticidal toxins, and these can be divided into insect immune suppressors via inhibition of phenoloxidase pathway:-1,2-benzene dicarboxylic acid (PA) (Ullah et al., 2014), benzylideneacetone (BZA) (Song et al., 2011), rhabduscin (Crawford et al., 2012; Eugenia Nuñez-Valdez et al., 2019), and 1,3-dihydroxy-2-(isopropyl)-5-(2-phenylethenyl)benzene (Eleftherianos et al., 2007); hemocyte pore-forming complexes: Xenorhabdus particulate toxins (Xpt) (Sheets et al., 2011), toxin complex toxins (Tc) (Blackburn et al., 1998), and Xenorhabdus α-xenorhabdolysin toxins (Xax) (Vigneux et al., 2007); apoptosis inducers: make caterpillar floppy toxins (Mcf) (Daborn et al., 2002; Dowling et al., 2004) and PaTox toxins (Jank et al., 2016); and those with yet unknown modes of action: PirAB (Yang et al., 2017) and xenocyloin (Proschak et al., 2014). Another ecological function of secreted metabolites is bioconversion by enzymes such as lipases, proteases, amylases, and proteases—their respective genes are enriched in Xenorhabdus and Photorhabdus genomes (Chaston et al., 2011)—creating a rich nutrient pool. To defend this from colonization by microbial competitors, a broad spectrum of antimicrobials is produced. These include antifungals: biocornutin (Böszörményi et al., 2009), cabanillasin (Houard et al., 2013), EP-19, GP-20 (Xiao et al., 2012), and xenocoumacin (Yang et al., 2011); antibacterials: darobactin (Imai et al., 2019), xenematide (Lang et al., 2008), photoditritide (Zhao et al., 2019), xenobactin (Grundmann et al., 2013), odilorhabdins (Pantel et al., 2018), xenorhabdin (McInerney et al., 1991), and PAX peptides (Gualtieri et al., 2009); antiprotozoals: phototemtide (Zhao et al., 2020), szentiamide (Ohlendorf et al., 2011), chaiyaphumins (Grundmann et al., 2014), rhabdopeptide/xenortide-like peptides (RXP) (Zhao L. et al., 2018), xenortide (Reimer et al., 2014), xenoamicin (Zhou et al., 2013), and ambactin (Schimming et al., 2014); and cytotoxic agents: fabclavines (Wenski et al., 2020) and phenylethylamine (PEA) derivatives (Proschak et al., 2011).

Recovered IJ3s leverage this nutrient-filled, enclosed environment to molt to fourth larval stage juvenile nematodes (J4) and then adults (Figure 1) that lay eggs after mating in the case of all steinernematids except Steinernema hermaphroditum—these species lay eggs without mating due to their hermaphroditic nature. This is like the androdioecious heterorhabditis whose adult females are also self-fertilized. Uniquely, Heterorhabditis adult females lay eggs into their uterus which hatch and develop into first larval stage juvenile nematodes (J1) through endotokia matricida (Clarke, 2020). Newly hatched EPNs molt from J1 through to J4 and then adults, which mate and lay eggs thus beginning another lifecycle. This continues until nutrients are depleted (Figure 1). Notably, infected cadavers are themselves protected from consumption by non-microbial competitors such as ants by the bacterial production of scavenger deterring factors (Zhou et al., 2002; Gulcu et al., 2012).

Upon nutrient depletion, J3 nematodes commence transformation to IJ3s by reassociating with bacterial symbionts (Figure 1)—this can be as few as one per nematode in the case of Xenorhabdus reassociations (Stilwell et al., 2018). Moreover, a highly species-specific reassociation occurs in Xenorhabdus-Steinernema complexes, and in Xenorhabdus nematophila, this is attributed to the NilC protein (Cowles and Goodrich-Blair, 2004). By retaining the second larval stage cuticle, J3s complete their transformation to IJ3s that then emigrate the cadaver in search of new insect prey. Notably, all seven macrocyclic antimicrobial non-ribosomal peptides (NRPs) with known toxicities—chaiyaphumins, photoditritide, szentiamide, xenobactin, phototemtide, xenoamicin, and PAX lipopeptides—were lowly toxic to mammalian cells—the lowest half-maximal inhibitory concentration (IC50) was 52 μM (Gualtieri et al., 2009; Ohlendorf et al., 2011; Grundmann et al., 2013, 2014; Zhou et al., 2013; Zhao et al., 2019, 2020). The bacteria possibly evolved to synthesize these compounds to inhibit diverse soil microorganisms while remaining lowly toxic to animal nematode hosts (Racine and Gualtieri, 2019). Biotechnologically, their low toxicity, natural derivatization, and macrocyclic structure (Dathe et al., 2004; Rodríguez et al., 2021) make them suitable for antibiotic drug development.

Xenorhabdus/Photorhabdus molecules in drug development

Many Xenorhabdus/Photorhabdus molecules have the potential to be developed into approved drugs (Challinor and Bode, 2015; Dreyer et al., 2018; Racine and Gualtieri, 2019; Booysen and Dicks, 2020). For example, Photorhabdus luminescens biosynthesized 3,5-dihydroxy-4-isopropylstilbene (Hu et al., 1997)—this is the active pharmaceutical ingredient in the drugs benvitimod and tapinarof (Zhang et al., 2022), which are approved for market in China and the USA, respectively, for the treatment of psoriasis and topical dermatitis (Lebwohl et al., 2021). NOS-502, an antibiotic lead compound currently in pre-clinical development, is a synthetic derivative of the odilorhabdins (Figure 1). These are cationic antimicrobial NRPs biosynthesized by X. nematophila that inhibit protein synthesis via unique sites on the 30S ribosome (Pantel et al., 2018). NOS-502 not only had a good in vivo safety profile but also inhibited beta-lactam resistant strains of both Escherichia coli and Klebsiella pneumoniae at minimum inhibitory concentrations (MICs) of 1.85 and 0.93 μM, respectively (Zhao M. et al., 2018). Another lead compound in pre-clinical development is darobactin A which is produced by Photorhabdus khanii (Lewis, 2020). It too was lowly toxic in murine models and inhibited beta-lactam resistant strains of both E. coli and K. pneumoniae at an MIC of 2.1 μM (Imai et al., 2019).

The development of novel antimalarial drugs is of current global health importance due to increasing resistance to artemisinin-based therapies in malaria-endemic regions such as East Africa (Asua et al., 2021), because of mutations in the Plasmodium falciparum K13 gene (Amaratunga et al., 2019). Two potential antimalarial lead compounds for pre-clinical development are the NRPs chaiyaphumin A from Xenorhabdus sp. PB61.4 (Grundmann et al., 2014) and rhabdopeptide/xenortide-like peptide (RXP) 6 from Xenorhabdus innexi (Zhao L. et al., 2018). This is because RXP 6 and chaiyaphumin A were inhibitory to P. falciparum at IC50 of 0.091 and 0.61 μM, respectively. Moreover, they had respective selectivity indexes of 63 and 151. Biochemically, the bioactivity of chaiyaphumin A was affected by the fatty acid acylated to its N terminal as the natural swapping of phenylacetic acid for n-butyrate created a derivative with an IC50 of 15.4 μM and selectivity index of 10. Thus, a probable route for creating chaiyaphumin derivatives with enhanced pharmacological properties is by swapping the Cstarter domain of its non-ribosomal peptide synthetase (NRPS) via NRPS re-engineering (Beck et al., 2020).

Although antibody–drug conjugates are promising anticancer therapies, their intrinsic high cost of development makes the price of approved drugs—such as enfortumab vedotin for the treatment of urothelial carcinoma—currently cost-ineffective (Wu et al., 2022). A possible solution is replacing the antibody component with a modified Photorhabdus Tc toxin, to translocate—within a cocoon-like structure (Roderer et al., 2019)—and deliver cytotoxic compounds into targeted cancer cells (Nǵanǵa et al., 2019). However, the concept that the Photorhabdus TcA subunit can selectively bind to a cancer cell needs to be first proven.

Conclusion

Twenty-seven Xenorhabdus, twenty-one Photorhabdus species, and four Serratia strains were identified as isolated from EPNs. Sixty-three novel species of Xenorhabdus could be discovered from corresponding characterized but under-investigated steinernematids. Due to the low phylogenetic diversity in the genus, the use of pangenome analyses for species delineation in Photorhabdus is recommended when enough genomes per species are available. The lifecycle of the nematode-bacterium complex is marked by the secretion of diverse bioactive bacterial molecules in the presence of juvenile nematodes, necessitating high selectivity. Thus, many of these molecules are applicable in biotechnology and a few are currently in drug development pipelines. This highlights the practical importance of discovering more nematophilic bacteria: They are a source of novel therapeutics.

Author contributions

RA did the research, wrote the manuscript, and approved the submitted version.